Molecular Detection of Foodborne Pathogens

MOLECULAR DETECTION OF FOODBORNE PATHOGENS

MOLECULAR DETECTION OF FOODBORNE PATHOGENS EDITED BY

DONGYOU LIU

Boca Raton London New York

CRC Press is an imprint of the Taylor & Francis Group, an informa business

CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2010 by Taylor and Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number: 978-1-4200-7643-1 (Hardback) This book contains information obtained from authentic and highly regarded sources. Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot assume responsibility for the validity of all materials or the consequences of their use. The authors and publishers have attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained. If any copyright material has not been acknowledged please write and let us know so we may rectify in any future reprint. Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data Molecular detection of foodborne pathogens / [edited by] Dongyou Liu. p. ; cm. Includes bibliographical references and index. ISBN 978-1-4200-7643-1 (hard back : alk. paper) 1. Foodborne diseases--Molecular diagnosis. 2. Food--Microbiology. I. Liu, Dongyou. II. Title. [DNLM: 1. Food Contamination--analysis. 2. Food Poisoning--microbiology. 3. Molecular Diagnostic Techniques--methods. WA 701 M718 2009] QR201.F62M65 2009 615.9’54--dc22 Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

2009017320

This book is dedicated to my parents, Jiaye Liu and Yunlian Li, whose unselfish sacrifice and unrelenting love have been a constant source of inspiration in my pursuit of knowledge and betterment.

Contents Preface........................................................................................................................................................................................xiii Editor........................................................................................................................................................................................... xv Contributors...............................................................................................................................................................................xvii Chapter 1. Molecular Detection: Principles and Methods......................................................................................................... 1 Lisa Gorski and Andrew Csordas

Section I  Foodborne Viruses Chapter 2. Adenoviruses.......................................................................................................................................................... 23 Charles P. Gerba and Roberto A. Rodríguez Chapter 3. Astroviruses........................................................................................................................................................... 33 Edina Meleg and Ferenc Jakab Chapter 4. Avian Influenza Virus............................................................................................................................................ 49 Giovanni Cattoli and Isabella Monne Chapter 5. Hepatitis A and E Viruses...................................................................................................................................... 63 Hiroshi Ushijima, Pattara Khamrin, and Niwat Maneekarn Chapter 6. Noroviruses............................................................................................................................................................ 75 Anna Charlotte Schultz, Jan Vinjé, and Birgit Nørrung Chapter 7. Rotaviruses............................................................................................................................................................. 91 Dongyou Liu, Larry A. Hanson, and Lesya M. Pinchuk Chapter 8. Sapoviruses...........................................................................................................................................................101 Grant S. Hansman Chapter 9. Slow Viral Diseases..............................................................................................................................................113 Takashi Onodera, Guangai Xue, Akikazu Sakudo, Gianluigi Zanusso, and Katsuaki Sugiura

Section II  Foodborne Gram-Positive Bacteria Chapter 10. Bacillus................................................................................................................................................................. 129 Noura Raddadi, Aurora Rizzi, Lorenzo Brusetti, Sara Borin, Isabella Tamagnini, and Daniele Daffonchio

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Chapter 11. Clostridium.......................................................................................................................................................... 145 Annamari Heikinheimo, Miia Lindström, Dongyou Liu, and Hannu Korkeala Chapter 12. Enterococcus........................................................................................................................................................ 157 Teresa Semedo-Lemsaddek, Rogério Tenreiro, Paula Lopes Alves, and Maria Teresa Barreto Crespo Chapter 13. Helicobacter..........................................................................................................................................................181 Norihisa Noguchi Chapter 14. Kocuria................................................................................................................................................................. 201 Edoardo Carretto and Daniela Barbarini Chapter 15. Listeria................................................................................................................................................................. 207 Dongyou Liu and Hans-Jürgen Busse Chapter 16. Micrococcus......................................................................................................................................................... 221 Friederike Hilbert and Hans-Jürgen Busse Chapter 17. Mycobacterium..................................................................................................................................................... 229 Irene R. Grant and Catherine E.D. Rees Chapter 18. Staphylococcus..................................................................................................................................................... 245 Paolo Moroni, Giuliano Pisoni, Paola Cremonesi, and Bianca Castiglioni Chapter 19. Streptococcus....................................................................................................................................................... 259 Mark van der Linden, Romney S. Haylett, Ralf René Reinert, and Lothar Rink

Section III  Foodborne Gram-Negative Bacteria Chapter 20. Aeromonas............................................................................................................................................................ 273 Germán Naharro, Jorge Riaño, Laura de Castro, Sonia Alvarez, and José María Luengo Chapter 21. Arcobacter............................................................................................................................................................ 289 Kurt Houf Chapter 22. Bacteriodes........................................................................................................................................................... 307 Rama Chaudhry, Anubhav Pandey, and Nidhi Sharma Chapter 23. Brucella.................................................................................................................................................................317 Sascha Al Dahouk, Karsten Nöckler, and Herbert Tomaso Chapter 24. Burkholderia.........................................................................................................................................................331 Karlene H. Lynch and Jonathan J. Dennis

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Chapter 25. Campylobacter..................................................................................................................................................... 345 Aurora Fernández Astorga and Rodrigo Alonso Chapter 26. Enterobacter......................................................................................................................................................... 361 Angelika Lehner, Roger Stephan, Carol Iversen, and Seamus Fanning Chapter 27. Escherichia........................................................................................................................................................... 369 Devendra H. Shah, Smriti Shringi, Thomas E. Besser, and Douglas R. Call Chapter 28. Klebsiella.............................................................................................................................................................. 391 Beatriz Meurer Moreira, Marco Antonio Lemos Miguel, Angela Christina Dias de Castro, Maria Silvana Alves, and Rubens Clayton da Silva Dias Chapter 29. Plesiomonas......................................................................................................................................................... 405 Jesús A. Santos, Andrés Otero, and María-Luisa García-López Chapter 30. Proteus..................................................................................................................................................................417 Antoni Róz∙alski and Paweł Sta˛ czek Chapter 31. Pseudomonas........................................................................................................................................................431 Olga Zaborina and John Alverdy Chapter 32. Salmonella............................................................................................................................................................ 447 Charlotta Löfström, Jeffrey Hoorfar, Jenny Schelin, Peter Rådström, and Burkhard Malorny Chapter 33. Serratia................................................................................................................................................................. 459 Zhi-Qing Hu, Wei-Hua Zhao, and Zhuting Hu Chapter 34. Shigella................................................................................................................................................................. 471 Benjamin R. Warren, Keith A. Lampel, and Keith R. Schneider Chapter 35. Vibrio.................................................................................................................................................................... 485 Asim K. Bej Chapter 36. Yersinia................................................................................................................................................................. 501 Mikael Skurnik, Peter Rådström, Rickard Knutsson, Bo Segerman, Saija Hallanvuo, Susanne Thisted Lambertz, Hannu Korkeala, and Maria Fredriksson-Ahomaa

Section IV  Foodborne Fungi Chapter 37. Alternaria............................................................................................................................................................. 521 Dongyou Liu, Stephen B. Pruett, and Cody Coyne Chapter 38. Aspergillus............................................................................................................................................................ 529 Giancarlo Perrone, Antonia Gallo, and Antonia Susca

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Chapter 39. Candida................................................................................................................................................................ 549 P. Lewis White, Samantha J. Hibbitts, Michael D. Perry, and Rosemary A. Barnes Chapter 40. Debaryomyces...................................................................................................................................................... 565 Juan J. Córdoba, Maria J. Andrade, Elena Bermúdez, Félix Núñez, Miguel A. Asensio, and Mar Rodríguez Chapter 41. Fusarium.............................................................................................................................................................. 577 Antonio Moretti and Antonia Susca Chapter 42. Penicillium........................................................................................................................................................... 593 Joëlle Dupont Chapter 43. Rhodotorula......................................................................................................................................................... 603 Diego Libkind and José Paulo Sampaio Chapter 44. Saccharomyces......................................................................................................................................................619 Franca Rossi and Sandra Torriani

Section V  Foodborne Protozoa Chapter 45. Acanthamoeba...................................................................................................................................................... 639 Hélène Yera, Pablo Goldschmidt, Christine Chaumeil, Muriel Cornet, and Marie-Laure Dardé Chapter 46. Cryptosporidium.................................................................................................................................................. 651 Una Ryan and Simone M. Cacciò Chapter 47. Cyclospora........................................................................................................................................................... 667 Dongyou Liu, G. Todd Pharr, and Frank W. Austin Chapter 48. Entamoeba........................................................................................................................................................... 677 Damien Stark and John Ellis Chapter 49. Encephalitozoon and Enterocytozoon................................................................................................................. 691 Jaco J. Verweij and Dongyou Liu Chapter 50. Giardia................................................................................................................................................................. 701 Yaoyu Feng and Lihua Xiao Chapter 51. Isospora.................................................................................................................................................................717 Somchai Jongwutiwes and Chaturong Putaporntip

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Chapter 52. Sarcocystis........................................................................................................................................................... 731 Benjamin M. Rosenthal Chapter 53. Toxoplasma...........................................................................................................................................................741 Chunlei Su and J.P. Dubey

Section VI  Foodborne Helminthes Chapter 54. Anisakis................................................................................................................................................................ 757 Stefano D’Amelio, Marina Busi, Sofia Ingrosso, Lia Paggi, and Elisabetta Giuffra Chapter 55. Clonorchis............................................................................................................................................................ 769 Heinz Mehlhorn, Boris Müller, and Jürgen Schmidt Chapter 56. Diphyllobothrium . .............................................................................................................................................. 781 Jean Dupouy-Camet and Hélène Yera Chapter 57. Fasciola................................................................................................................................................................ 789 Xing-Quan Zhu, Qing-Jun Zhuang, Rui-Qing Lin, and Wei-Yi Huang Chapter 58. Heterophyidae...................................................................................................................................................... 795 Ron Dzikowski and Michael G. Levy Chapter 59. Metagonimus........................................................................................................................................................ 805 Jae-Ran Yu and Jong-Yil Chai Chapter 60. Opisthorchis......................................................................................................................................................... 813 Paiboon Sithithaworn, Thewarach Laha, and Ross H. Andrews Chapter 61. Paragonimus........................................................................................................................................................ 827 Kanwar Narain, Takeshi Agatsuma, and David Blair Chapter 62. Taenia................................................................................................................................................................... 839 Akira Ito, Minoru Nakao, Yasuhito Sako, Kazuhiro Nakaya, Tetsuya Yanagida, and Munehiro Okamoto Chapter 63. Trichinella............................................................................................................................................................ 851 Edoardo Pozio and Giuseppe La Rosa Index.......................................................................................................................................................................................... 865

Preface Foodborne pathogens are microorganisms (e.g., bacteria, viruses, fungi, and parasites) that are capable of infecting humans via contaminated food and/or water. In recent years, diseases caused by foodborne pathogens have become an important public health problem worldwide, resulting in significant morbidity and mortality. Currently, there are over 250 known foodborne diseases. Due to the introduction of pathogens to other geographic regions through population movement and globalization of the food supply, new foodborne infections are continuously emerging. Furthermore, pathogen evolution, changes in human immune status and life-style as well as food manufacturing practices also contribute to increased incidences of foodborne ­illnesses. As a consequence, large outbreaks of foodborne diseases have been reported with alarming frequencies. It is well known that one of the most effective ways to control and prevent human foodborne infections is to implement a surveillance system that includes a capability to rapidly and precisely detect, identify, and monitor foodborne pathogens at the nucleic acid level. The purpose of this book is to bring out an all-encompassing volume on the detection and identification of major foodborne bacterial, fungal, viral, and parasitic pathogens using state-of-art molecular techniques. Each chapter includes a concise review of the pathogen concerned with respect to its biology, epidemiology, and pathogenesis; a summary of the molecular detection methods available; a description of clinical/food sample collection and preparation procedures; a selection of robust, effective, step-wise molecular detection protocols for each pathogen; and a discussion on the challenges and continuing research needs to further extend the utility and performance of molecular diagnostic methods for foodborne diseases. With each chapter written by scientists with expertise in their respective foodborne pathogen research, this book provides comprehensive coverage of the molecular methodologies for the detection and identification of major foodborne pathogens. It is an indispensable tool for clinical, food, and industrial laboratory scientists involved in the diagnosis of foodborne diseases; a convenient textbook for prospective undergraduate and graduate students intending to pursue a career in food microbiology and medical technology; and a reliable reference for upcoming and experienced laboratory scientists wishing to develop and polish their skills in the molecular detection of major foodborne pathogens. Given the number of foodborne pathogens covered, and the breadth and depth of the topics discussed, an inclusive book like this is undoubtedly beyond the capacity of an individual’s effort. It is my fortune and honor to have a large panel of international scientists as chapter contributors, whose willingness to share their technical insights on foodborne pathogen detection has made this book possible. Moreover, the professionalism and dedication of senior editor, Steve Zollo, and other editorial staff at CRC Press have contributed to its enhanced presentation. I hope the readers will find it as stimulating and rewarding as I do through reading this book, which by presenting relevant background information and ready-to-run molecular detection protocols will serve to save readers’ time and patients’ lives. Dongyou Liu, PhD

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Editor Dongyou Liu, PhD, is currently a member of the research faculty in the Department of Basic Sciences, College of Veterinary Medicine at Mississippi State University in Starkville. In 1982, he graduated with a veterinary science degree from Hunan Agricultural University in China. After one year of postgraduate training under the supervision of Professor Kong Fangyao at Beijing Agricultural University (presently China Agricultural University) in China, he completed his PhD study on the immunological diagnosis of human hydatid disease due to the parasitic tapeworm Echinococcus granulosus in the laboratory of Drs. Michael D. Rickard and Marshall W. Lightowlers at the University of Melbourne School of Veterinary Science in Australia in 1989. During the past two decades, he has worked in several research and clinical laboratories in Australia and the United States, with an emphasis on molecular microbiology, especially in the development of nucleic acid-based assays for species- and virulence-specific determination of microbial pathogens such as ovine footrot bacterium (Dichelobacter nodosus), dermatophyte fungi (Trichophyton, Microsporum, and Epidermophyton), and listeriae (Listeria species). He is the editor of the Handbook of Listeria monocytogenes and the Handbook of Nucleic Acid Purification, both of which have been published recently by Taylor & Francis/CRC Press.

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Contributors Takeshi Agatsuma Department of Environmental Health Sciences Kochi Medical School Nankoku City, Kochi, Japan Rodrigo Alonso Departamento de Inmunología, Microbiología y Parasitología Facultad de Farmacia Universidad del Pais Vasco/Euskal Herriko Unibertsitatea Vitoria-Gasteiz, Spain Sonia Alvarez Department of Animal Health University of León León, Spain John Alverdy Center for Surgical Infection Research and Therapeutics University of Chicago Chicago, Illinois Maria Silvana Alves Faculdade de Farmácia e Bioquímica Universidade Federal de Juiz de Fora Minas Gerais, Brazil Paula Lopes Alves Instituto de Biologia Experimental e Tecnológica (IBET) Av. da República, Quinta do Marquês Oeiras, Portugal M.J. Andrade Higiene y Seguridad Alimentaria Facultad de Veterinaria Universidad de Extremadura Cáceres, Spain Ross H. Andrews School of Pharmacy and Medical Sciences University of South Australia Adelaide, Australia M.A. Asensio Higiene y Seguridad Alimentaria Facultad de Veterinaria Universidad de Extremadura Cáceres, Spain

Aurora Fernández Astorga Departamento de Inmunología, Microbiología y Parasitología Facultad de Farmacia Universidad del Pais Vasco/Euskal Herriko Unibertsitatea Vitoria-Gasteiz, Spain Frank W. Austin Department of Basic Sciences College of Veterinary Medicine Mississippi State University Mississippi State, Mississippi Daniela Barbarini Bacteriology Laboratory Infectious Diseases, Laboratories of Experimental Researches Fondazione “IRCCS Policlinico San Matteo” Pavia, Italy Rosemary A. Barnes Department of Medical Microbiology Cardiff University University Hospital of Wales Cardiff, Wales, United Kingdom Asim K. Bej Department of Biology University of Alabama at Birmingham Birmingham, Alabama E. Bermúdez Higiene y Seguridad Alimentaria Facultad de Veterinaria Universidad de Extremadura Cáceres, Spain Thomas E. Besser Department of Veterinary Microbiology and Pathology College of Veterinary Medicine Washington State University Pullman, Washington David Blair School of Marine and Tropical Biology James Cook University Townsville, Australia Sara Borin Department of Food Science and Microbiology University of Milan Milan, Italy xvii

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Contributors

Lorenzo Brusetti Department of Food Science and Microbiology University of Milan Milan, Italy

Jong-Yil Chai Department of Parasitology and Tropical Medicine Seoul National University College of Medicine Seoul, Korea

Marina Busi Department of Public Health Science Sapienza University of Rome Rome, Italy

Rama Chaudhry Department of Microbiology All India Institute of Medical Sciences New Delhi, India

Hans-Jürgen Busse Institute of Bacteriology, Mycology and Hygiene University of Veterinary Medicine Vienna, Austria

Christine Chaumeil Laboratoire du Centre National d’Ophtalmologie des Quinze-Vingts Paris, France

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

J.J. Córdoba Higiene y Seguridad Alimentaria Facultad de Veterinaria Universidad de Extremadura Cáceres, Spain

Douglas R. Call Department of Veterinary Microbiology and Pathology College of Veterinary Medicine Washington State University Pullman, Washington

Muriel Cornet Laboratoire de Microbiologie Hôpital Hôtel-Dieu Paris, France

Edoardo Carretto Bacteriology Laboratory Infectious Diseases, Laboratories of Experimental Researches Fondazione “IRCCS Policlinico San Matteo” Pavia, Italy Bianca Castiglioni Institute of Agricultural Biology and Biotechnology Italian National Research Council Milan, Italy Angela Christina Dias de Castro Instituto de Microbiologia Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Laura de Castro Department of Animal Health University of León León, Spain Giovanni Cattoli Istituto Zooprofilattico Sperimentale delle Venezie Research and Development Department OIE/FAO and National Reference Laboratory for Newcastle Disease and Avian Influenza OIE Collaborating Center for Epidemiology, Training and Control of Emerging Avian Diseases Legnaro, Padova, Italy

Cody Coyne Department of Basic Sciences College of Veterinary Medicine Mississippi State University Mississippi State, Mississippi Paola Cremonesi Institute of Agricultural Biology and Biotechnology Italian National Research Council Milan, Italy Maria Teresa Barreto Crespo Instituto de Biologia Experimental e Tecnológica (IBET) Av. da República, Quinta do Marquês Oeiras, Portugal Andrew Csordas Institute for Collaborative Biotechnologies University of California, Santa Barbara Santa Barbara, California Daniele Daffonchio Department of Food Science and Microbiology University of Milan Milan, Italy Sascha Al Dahouk Department of Internal Medicine III RWTH Aachen University Aachen, Germany

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Contributors

Stefano D’Amelio Department of Public Health Science Sapienza University of Rome Rome, Italy

Yaoyu Feng School of Resource and Environmental Engineering East China University of Science and Technology Shanghai, People’s Republic of China

Marie-Laure Dardé Laboratoire de Parasitologie-Mycologie CHU Limoges, France

Maria Fredriksson-Ahomaa Institute of Hygiene and Technology of Food of Animal Origin Ludwig-Maximilian University Munich, Germany

Jonathan J. Dennis Department of Biological Sciences University of Alberta Edmonton, Alberta, Canada Rubens Clayton da Silva Dias Division of Infectious Diseases and Immunity School of Public Health University of California Berkeley, California J.P. Dubey United States Department of Agriculture, Agricultural Research Service Animal and Natural Resources Institute Animal Parasitic Diseases Laboratory Beltsville, Maryland Joëlle Dupont Muséum National d’Histoire Naturelle Département Systématique et Evolution Paris, France Jean Dupouy-Camet Laboratoire de Parasitologie-Mycologie Hôpital Cochin AP-H Université Paris Descartes Paris, France Ron Dzikowski Department of Microbiology & Molecular Genetics The Kuvin Center for the Study of Infectious and Tropical Diseases The Institute for Medical Research Israel-Canada The Hebrew University–Hadassah Medical School Jerusalem, Israel John Ellis Department of Medical and Molecular Biosciences University of Technology Sydney Broadway, Australia Seamus Fanning Centre for Food Safety, School of Agriculture, Food Science and Veterinary Medicine Veterinary Sciences Centre University College Dublin Dublin, Ireland

Antonia Gallo Institute of Sciences of Food Production National Research Council (ISPA-CNR) Bari, Italy María-Luisa García-López Department of Food Hygiene and Food Microbiology University of León León, Spain Charles P. Gerba Department of Soil, Water and Environmental Science University of Arizona Tucson, Arizona Elisabetta Giuffra Parco Tecnologico Padano Lodi, Italy Pablo Goldschmidt Laboratoire du Centre National d’Ophtalmologie des Quinze-Vingts Paris, France Lisa Gorski Produce Safety and Microbiology Research Unit United States Department of Agriculture Agricultural Research Service Western Regional Research Center Albany, California Irene R. Grant School of Biological Sciences Queen’s University Belfast Belfast, Northern Ireland, United Kingdom Saija Hallanvuo Department of Animal Diseases and Food Safety Research Finnish Food Safety Authority Evira Helsinki, Finland Grant S. Hansman Department of Virology II National Institute of Infectious Diseases Musashi-murayama Tokyo, Japan

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Larry A. Hanson Department of Basic Sciences College of Veterinary Medicine Mississippi State University Mississippi State, Mississippi Romney S. Haylett Institute of Immunology RWTH Aachen University Hospital Aachen, Germany Annamari Heikinheimo Department of Food and Environmental Hygiene University of Helsinki Helsinki, Finland Samantha J. Hibbitts Department of Obstetrics and Gynaecology Cardiff University University Hospital of Wales Cardiff, Wales, United Kingdom Friederike Hilbert Institute of Meat Hygiene, Meat Technology and Food Science University of Veterinary Medicine Vienna, Austria Jeffrey Hoorfar National Food Institute Technical University of Denmark Søborg, Denmark Kurt Houf Department of Veterinary Public Health and Food Safety Ghent University Merelbeke, Belgium Zhi-Qing Hu Department of Microbiology and Immunology Showa University School of Medicine Tokyo, Japan Zhuting Hu Department of Biology International Christian University Tokyo, Japan

Contributors

Akira Ito Department of Parasitology Asahikawa Medical College Asahikawa, Japan Carol Iversen Centre for Food Safety, School of Agriculture, Food Science and Veterinary Medicine Veterinary Sciences Centre University College Dublin Dublin, Ireland Ferenc Jakab Department of Genetics and Molecular Biology Institute of Biology, Faculty of Sciences University of Pécs Pécs, Hungary Somchai Jongwutiwes Molecular Biology of Malaria and Opportunistic Parasites Research Unit Department of Parasitology Chulalongkorn University Bangkok, Thailand Pattara Khamrin Aino Health Science Center Aino University Tokyo, Japan Rickard Knutsson Department of Bacteriology National Veterinary Institute Uppsala, Sweden Hannu Korkeala Department of Food and Environmental Hygiene University of Helsinki Helsinki, Finland Thewarach Laha Department of Parasitology Liver Fluke and Cholangiocarcinoma Research Center Khon Kaen University Khon Kaen, Thailand

Wei-Yi Huang Department of Veterinary Medicine College of Animal Science and Technology Guangxi University Nanning, Guangxi, People’s Republic of China

Susanne Thisted Lambertz Research and Development Department National Food Administration Uppsala, Sweden

Sofia Ingrosso Department of Public Health Science Sapienza University of Rome Rome, Italy

Keith A. Lampel Food and Drug Administration Division of Microbiology College Park, Maryland

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Contributors

Angelika Lehner Institute for Food Safety and Hygiene University of Zurich Zurich, Switzerland

Burkhard Malorny Federal Institute for Risk Assessment National Salmonella Reference Laboratory Berlin, Germany

Michael G. Levy Department of Population Health and Pathobiology College of Veterinary Medicine North Carolina State University Raleigh, North Carolina

Niwat Maneekarn Department of Microbiology, Chiang Mai University Chiang Mai, Thailand

Diego Libkind Laboratorio de Microbiología Aplicada y Biotecnología Instituto de Investigaciones en Biodiversidad y Medio Ambiente (INIBIOMA) Universidad Nacional del Comahue CRUB – Consejo Nacional de Investigaciones Científicas y Tecnológicas (CONICET) Bariloche, Río Negro, Argentina Rui-Qing Lin Laboratory of Parasitology College of Veterinary Medicine South China Agricultural University Guangzhou, Guangdong, People’s Republic of China Mark van der Linden Institute of Medical Microbiology and National Reference Center for Streptococci RWTH Aachen University Hospital Aachen, Germany Miia Lindström Department of Food and Environmental Hygiene University of Helsinki Helsinki, Finland Dongyou Liu Department of Basic Sciences College of Veterinary Medicine Mississippi State University Mississippi State, Mississippi Charlotta Löfström National Food Institute Technical University of Denmark Søborg, Denmark

Heinz Mehlhorn Department of Parasitology Heinrich Heine University Düsseldorf, Germany Edina Meleg Department of Biophysics Faculty of Medicine University of Pécs Pécs, Hungary Marco Antonio Lemos Miguel Instituto de Microbiologia Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Isabella Monne Istituto Zooprofilattico Sperimentale delle Venezie Research and Development Department OIE/FAO and National Reference Laboratory for Newcastle Disease and Avian Influenza OIE Collaborating Center for Epidemiology, Training and Control of Emerging Avian Diseases Legnaro, Padova, Italy Beatriz Meurer Moreira Instituto de Microbiologia Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Antonio Moretti Institute of Sciences of Food Production National Research Council (ISPA-CNR) Bari, Italy

José María Luengo Department of Biochemistry and Molecular Biology University of León León, Spain

Paolo Moroni Department of Veterinary Pathology Hygiene and Public Health University of Milan Milan, Italy

Karlene H. Lynch Department of Biological Sciences University of Alberta Edmonton, Alberta, Canada

Boris Müller Department of Parasitology Heinrich Heine University Düsseldorf, Germany

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Contributors

Germán Naharro Department of Animal Health University of León León, Spain

Lia Paggi Department of Public Health Science Sapienza University of Rome Rome, Italy

Minoru Nakao Department of Parasitology Asahikawa Medical College Asahikawa, Japan

Anubhav Pandey Department of Microbiology All India Institute of Medical Sciences New Delhi, India

Kazuhiro Nakaya Laboratory Animals for Medical Research Asahikawa Medical College Asahikawa, Japan

Giancarlo Perrone Institute of Sciences of Food Production National Research Council (ISPA-CNR) Bari, Italy

Kanwar Narain Regional Medical Research Centre, N.E. Region Indian Council of Medical Research Dibrugarh, Assam, India

Michael D. Perry NPHS Microbiology Cardiff University Hospital of Wales Cardiff, Wales, United Kingdom

Karsten Nöckler Federal Institute for Risk Assessment Berlin, Germany Norihisa Noguchi Department of Microbiology School of Pharmacy Tokyo University of Pharmacy and Life Sciences Hachioji, Tokyo, Japan Birgit Nørrung Department of Veterinary Pathobiology University of Copenhagen Frederiksberg C, Denmark F. Nuñez Higiene y Seguridad Alimentaria Facultad de Veterinaria Universidad de Extremadura Cáceres, Spain Munehiro Okamoto Department of Parasitology School of Veterinary Medicine Tottori University Tottori, Japan Takashi Onodera Department of Molecular Immunology, School of Agricultural and Life Sciences University of Tokyo Bunkyo-ku, Tokyo, Japan Andrés Otero Department of Food Hygiene and Food Microbiology University of León León, Spain

G. Todd Pharr Department of Basic Sciences College of Veterinary Medicine, Mississippi State University Mississippi State, Mississippi Lesya M. Pinchuk Department of Basic Sciences College of Veterinary Medicine Mississippi State University Mississippi State, Mississippi Giuliano Pisoni Department of Veterinary Pathology Hygiene and Public Health University of Milan Milan, Italy Edoardo Pozio Department of Infectious, Parasitic and Immunomediated Diseases Istituto Superiore di Sanità Rome, Italy Stephen B. Pruett Department of Basic Sciences College of Veterinary Medicine, Mississippi State University Mississippi State, Mississippi Chaturong Putaporntip Molecular Biology of Malaria and Opportunistic Parasites Research Unit Department of Parasitology Chulalongkorn University Bangkok, Thailand

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Contributors

Noura Raddadi Department of Food Science and Microbiology University of Milan Milan, Italy

Franca Rossi Dipartimento di Biotecnologie Universita degli Studi di Verona Verona, Italy

Peter Rådström Applied Microbiology Lund Institute of Technology Lund University Lund, Sweden

Antoni Róz∙ alski Institute of Microbiology and Immunology University of Łódz´ Łódz´, Poland

Catherine E.D. Rees School of Biosciences University of Nottingham Sutton Bonington Campus Leicestershire, England, United Kingdom Ralf René Reinert Wyeth Vaccines Research Paris la Défense, France Jorge Riaño Department of Animal Health University of León León, Spain Lothar Rink Institute of Immunology RWTH Aachen University Hospital Aachen, Germany Aurora Rizzi Department of Food Science and Microbiology University of Milan Milan, Italy M. Rodríguez Higiene y Seguridad Alimentaria Facultad de Veterinaria Universidad de Extremadura Cáceres, Spain Roberto A. Rodríguez Department of Environmental Science and Engineering University of North Carolina Chapel Hill, North Carolina Giuseppe La Rosa Department of Infectious, Parasitic and Immunomediated Diseases Istituto Superiore di Sanità Rome, Italy Benjamin M. Rosenthal United States Department of Agriculture Agricultural Research Service Animal Natural Resources Institute Animal Parasitic Diseases Laboratory Beltsville, Maryland

Una Ryan Division of Health Sciences School of Veterinary and Biomedical Science Murdoch University Perth, Australia Yasuhito Sako Department of Parasitology Asahikawa Medical College Asahikawa, Japan Akikazu Sakudo Department of Virology, Research Institute for Microbial Diseases Osaka University Suita, Osaka, Japan José Paulo Sampaio Centro de Recursos Microbiológicos Departamento de Ciências da Vida Universidade Nova de Lisboa Caparica, Portugal Jesús A. Santos Department of Food Hygiene and Food Microbiology University of León León, Spain Jenny Schelin Applied Microbiology Lund Institute of Technology Lund University Lund, Sweden Jürgen Schmidt Department of Parasitology Heinrich Heine University Düsseldorf, Germany Keith R. Schneider Food Science and Human Nutrition Department University of Florida Gainesville, Florida Anna Charlotte Schultz National Food Institute Technical University of Denmark (DTU) Søborg, Denmark

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Contributors

Bo Segerman Department of Bacteriology National Veterinary Institute Uppsala, Sweden

Roger Stephan Institute for Food Safety and Hygiene University of Zurich Zurich, Switzerland

Teresa Semedo-Lemsaddek Universidade de Lisboa Center for Biodiversity Functional and Integrative Genomics (BioFIG) Edifício ICAT, Campus da FCUL, Campo Grande Lisbon, Portugal

Chunlei Su Department of Microbiology University of Tennessee Knoxville, Tennessee

Devendra H. Shah Department of Veterinary Microbiology and Pathology College of Veterinary Medicine Washington State University Pullman, Washington Nidhi Sharma Department of Microbiology All India Institute of Medical Sciences New Delhi, India Smriti Shringi Department of Veterinary Microbiology and Pathology College of Veterinary Medicine Washington State University Pullman, Washington Paiboon Sithithaworn Department of Parasitology Liver Fluke and Cholangiocarcinoma Research Center Khon Kaen University Khon Kaen, Thailand Mikael Skurnik Department of Bacteriology and Immunology Infection Biology Research Program, Haartman Institute, University of Helsinki Helsinki, Finland and Helsinki University Central Hospital Laboratory Diagnostics Helsinki, Finland

Katsuaki Sugiura Food and Agricultural Materials Inspection Centre Chuo-ku, Saitama-shi Saitama, Japan Antonia Susca Institute of Sciences of Food Production National Research Council (ISPA-CNR) Bari, Italy Isabella Tamagnini Department of Food Science and Microbiology University of Milan Milan, Italy Rogério Tenreiro Universidade de Lisboa Center for Biodiversity Functional and Integrative Genomics (BioFIG) Edifício ICAT, Campus da FCUL, Campo Grande Lisbon, Portugal Herbert Tomaso Friedrich Loeffler Institute Institute of Bacterial Infections and Zoonoses Jena, Germany Sandra Torriani Dipartimento di Scienze, Tecnologie e Mercati della Vite e del Vino Università degli Studi di Verona Verona, Italy

Paweł Sta˛ czek Institute of Microbiology and Immunology University of Łódz´ Łódz´, Poland

Hiroshi Ushijima Aino Health Science Center Aino University Tokyo, Japan

Damien Stark Division of Microbiology, SydPath St. Vincent‘s Hospital Darlinghurst, Australia

Jaco J. Verweij Department of Parasitology Leiden University Medical Center Leiden, the Netherlands

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Contributors

Jan Vinjé Division of Viral Diseases Center for Disease Control (CDC) Atlanta, Georgia

Jae-Ran Yu Department of Environmental and Tropical Medicine Konkuk University School of Medicine Seoul, Korea

Benjamin R. Warren Research, Quality, & Innovation ConAgra Foods, Inc. Omaha, Nebraska

Olga Zaborina Center for Surgical Infection Research and Therapeutics University of Chicago Chicago, Illinois

P. Lewis White NPHS Microbiology Cardiff University Hospital of Wales Cardiff, Wales, United Kingdom Lihua Xiao Division of Parasitic Diseases Centers for Disease Control and Prevention Atlanta, Georgia Guangai Xue Department of Molecular Immunology School of Agricultural and Life Sciences University of Tokyo Bunkyo-ku, Tokyo, Japan Tetsuya Yanagida Department of Parasitology Asahikawa Medical College Asahikawa, Japan Hélène Yera Laboratoire de Parasitologie-Mycologie Hôpital Cochin AP-HP Université Paris Descartes Paris, France

Gianluigi Zanusso Department of Neurological Sciences University of Verona Verona, Italy Wei-Hua Zhao Department of Microbiology and Immunology Showa University School of Medicine Tokyo, Japan Xing-Quan Zhu Laboratory of Parasitology College of Veterinary Medicine South China Agricultural University Guangzhou, Guangdong, People’s Republic of China Qing-Jun Zhuang Laboratory of Parasitology College of Veterinary Medicine South China Agricultural University Guangzhou, Guangdong, People’s Republic of China

Detection: 1 Molecular Principles and Methods Lisa Gorski

United States Department of Agriculture

Andrew Csordas

University of California

Contents 1.1 Introduction........................................................................................................................................................................... 1 1.2 Detection Methods................................................................................................................................................................ 3 1.2.1 Pathogen Detection in Complex Matrices—Sample Preparation.............................................................................. 3 1.2.2 Nucleic Acid Based Detection................................................................................................................................... 3 1.2.2.1 PCR............................................................................................................................................................. 3 1.2.2.2 Isothermal Amplification............................................................................................................................ 7 1.2.2.3 Microarray Detection.................................................................................................................................. 8 1.2.3 Fluorescence in situ Hybridization (FISH)................................................................................................................ 8 1.2.4 Immunological Detection Methods........................................................................................................................... 8 1.2.5 Combined Detection Methods................................................................................................................................... 9 1.2.6 Foodborne Pathogen Typing...................................................................................................................................... 9 1.2.7 Microfabrication and Microfluidics........................................................................................................................... 9 1.2.8 Other Molecular Detection Approaches.................................................................................................................... 9 1.2.9 Assay Design and Data Analysis Software.............................................................................................................. 10 1.3 Detection Targets................................................................................................................................................................. 10 1.3.1 Viral Targets............................................................................................................................................................ 10 1.3.1.1 RNA Targets.............................................................................................................................................. 11 1.3.1.2 Viral Structural Genes.............................................................................................................................. 11 1.3.1.3 Other Viral Targets.................................................................................................................................... 11 1.3.2 Nonviral Targets...................................................................................................................................................... 11 1.3.2.1 Ribosomal RNA Genes . .......................................................................................................................... 11 1.3.2.2 Cytoskeleton Proteins................................................................................................................................ 12 1.3.2.3 Virulence and Toxin Genes....................................................................................................................... 12 1.3.2.4 Unique Genes and Sequences................................................................................................................... 12 1.3.2.5 Insertion Elements..................................................................................................................................... 13 1.3.2.6 Mitochondrial Genes................................................................................................................................. 13 1.3.2.7 Genes for Surface Expressed Markers...................................................................................................... 13 1.3.3 Using Multiple Targets............................................................................................................................................. 14 1.4 Validation............................................................................................................................................................................ 14 1.5 Conclusions.......................................................................................................................................................................... 14 Acknowledgments........................................................................................................................................................................ 15 References.................................................................................................................................................................................... 15

1.1  Introduction While the vast majority of our food supplies are nutritious and safe, illness due to foodborne pathogens still affects millions if not billions of people each year. It is estimated that up to 30% of the population in industrialized nations suffer

from foodborne illness each year.1 In the U.S. there are an estimated 76 million cases each year that result in 325,000 hospitalizations, and 5000 deaths.2 Estimates of the number of cases in developing countries are difficult to obtain due to differences in reporting of cases in different countries; however, the rates of illness are expected to be higher.1,3,4 1

2

Molecular Detection of Foodborne Pathogens

Diarrheal diseases, a high number of which result from foodborne contamination, kill an estimated 1.8 million children worldwide.3 Table 1.1 summarizes the statistics of U.S. foodborne illness outbreaks for the year 2006 broken down by etiology. An outbreak is constituted by more than one person becoming ill by the same strain of an organism. The list displays only outbreaks from known etiologies of bacterial, viral, parasitic, and helminthic origin, and does not take into account outbreaks where an etiology could not be assigned. Nor does it take into account sporadic cases of illness, which far outnumber outbreak cases. Most of these sporadic cases are not reported to any official health tracking agency because they are not severe, or cultures are never obtained.1 An even greater number of people with sporadic cases of foodborne illness do not seek medical attention. Whether an illness is mild or severe, the underlying message from the statistics is that millions or billions of servings of food are contaminated with a pathogen or a toxin each year. Table 1.1 illustrates that the types of foods implicated is broad and comprises meats, dairy, produce, grains, processed foods, and water. While many cases of foodborne illness result from human cross-contamination in restaurants or in the home, a large amount results from foods that arrive

into the kitchen already contaminated. These organisms can contaminate the foods directly by association with feed animals or plants prior to or during processing, through contaminated water used for watering or washing, and through handling by infected people. One of the most difficult and fundamental issues in food safety is the detection of foodborne pathogens. The problem is terribly complex with a multitude of factors and variables with which to contend. With the infectious dose of some of the pathogens as low as <100 cells or particles, sensitivity is essential. In some instances, an enrichment step is necessary to amplify the number of pathogens in the sample simply so that they can be detected. However, enrichment does not work with viruses or toxins, and some organisms with long generation times can take weeks to enrich. Additionally nonprocessed or minimally processed foods are not sterile and native microflora can sometimes mask the presence of the pathogen. Finally the food matrix itself sometimes inhibits detection by affecting the chemistries used in detection methods. While an all-encompassing test that would detect every possible pathogen or toxin would be desirable, the technology does not yet exist. Ideally, the detection of pathogens should be fast and economical. Ultimately a balance between the financial burden of testing and the risk of selling of untested foods must

Table 1.1 Number of Foodborne Outbreaks with Confirmed Etiologies in the U.S. for the Year 2006 Agent

No. of Outbreaks

No. of Cases

Bacterial

Bacillus Brucella Campylobacter Clostridium Escherichia Listeria Salmonella Shigella Staphylococcus Vibrio Subtotal

3 1 22 20 29 3 116 9 12 8 223

35 5 283 745 520 7 2751 183 380 427 5336

Viral

Hepatitis A Norovirus Subtotal

3 333 336

34 10,970 11,004

Parasite

Cryptosporidium Cyclospora Giardia Subtotal

2 3 2 7

16 19 11 46

Helminth

Trichinella Subtotal

1 1

2 2

567

16,388

Total

Suspected Vehicles Produce, rice, meat Cheese Milk, cheese, seafood, produce, meat Produce, seafood, canned food, meat Milk, produce, meat Cheese, salad Meat, dairy, produce, peanut butter Salad, produce, meat Meat, dairy, seafood Seafood

Spring water, unspecified Salads, seafood, meat, produce Unspecified Fruit salad Unspecified Bear meat

Source: Compiled from Centers for Disease Control http://www.cdc.gov.

3

Molecular Detection: Principles and Methods

be met to ensure the safety of consumers and simultaneous profitability for food producers. The following chapters give detailed reviews of the latest methods and targets for detection of specific organisms. However, when reviewing the subject of molecular detection methods, common themes arise. These themes relate to the choices of detection methods and the molecular targets for detection.

1.2 Detection Methods A wide range of foodborne pathogen detection techniques have been developed including culturing methods, nucleic acid methods, immunological methods, microscopy, spectroscopy, and bioluminescence, with varying degrees of cost, specificity, sensitivity, and ease of use. The major considerations of a detection system include the cost of the process, the target for detection, and the specificity and sensitivity of the procedure selected for detection. In recent years painstaking methods of cell culture and microscopic observation have yielded faster, more efficient molecular methods of detection. While traditional microbial detection methods may yield adequate target specificity and sensitivity, the time to results is on the order of days, often relying upon pathogen growth. Numerous molecular techniques have emerged that offer the advantage of speed along with specific and sensitive detection. Molecular methods have also proven advantageous in cases where it is difficult to culture the target of interest, as can be the case with viruses. These methods require a solid understanding of the physiology of the target organism, its close relatives, and those with which it may coexist on a food surface.

1.2.1 Pathogen Detection in Complex Matrices—Sample Preparation Simultaneous advances in detection methods and in sample preparation prior to analysis are needed to ensure a safe food supply.5 Foodborne pathogens have been associated with a wide variety of foods including poultry, beef, shellfish, fruits, vegetables, and drinking water. Without appropriate preparation of a test sample prior to detection, a common potential problem to many detection methods is that the sample background material may drastically decrease the sensitivity of the detection step or even lead to false negative test results. Food derived polymerase chain reaction (PCR) inhibitors include Ca2+, fats, glycogen, and phenolic compounds.6 The presence of proteinases in cheese7 and milk8 may also inhibit PCR. Different approaches have been used to counteract poor PCR performance in difficult backgrounds. Bovine serum albumin has shown success in relieving PCR inhibition in certain cases,9 and the type of DNA polymerase used can greatly affect the outcome of a reaction in the presence of biological samples.10 Additional potential challenges of detecting foodborne pathogens include their nonuniform dispersal and very low concentrations within foods. Therefore, considerable effort is often required to prepare a sample such that it is suitable

for testing with a nucleic acid detection procedure. Methods that have been used for the removal of PCR inhibitors include physical separation techniques such as filtration, DNA extraction, and adsorptive methods such as immunomagnetic separation.11 Lampel et al.12 used filters capable of trapping and lysing microorganisms, then used these filters directly in PCR reactions. The detection limits of Shigella flexneri in artificially contaminated foods using the filter system were greatly improved as compared to unfiltered tests.

1.2.2 Nucleic Acid Based Detection Advances in nucleic acid testing have included rapid amplification techniques and associated automated instrumentation, microarray based technology, and lab-on-a-chip platforms. The relatively low cost and speed of oligonucleotide synthesis, the wide range of 3′ and 5′ oligonucleotide modifications readily available, and powerful software to aid in molecular assay design and data analysis have facilitated the growth of a wide range of nucleic acid based techniques applied to the detection of foodborne pathogens. 1.2.2.1  PCR Nucleic acid amplification techniques have an enormous range of applications and have become an indispensible tool in molecular biology and powerful rapid screening method in the detection of foodborne pathogens. By targeting and amplifying (or making copies of) DNA sequences in vitro, it has been possible to detect the presence of specific DNA sequences with sensitivities down to a single target copy per reaction, and in many cases quantify the results. PCR is a method for the amplification of double or single stranded (ss) DNA sequences in vitro. The reaction proceeds in response to temperature driven steps of double stranded (ds) DNA denaturation, primer or ss oligonucleotide annealing to complementary ss target DNA sequences, and DNA polymerase extension. These steps are repeated, and under appropriate conditions will generate a doubling of the initial number of target copy sequences with each cycle. The primers define the 5′ ends of the discrete products that are subsequently formed. Three step PCRs use three individual temperature steps for denaturation, annealing, and extension, while two step PCRs use a combined annealing and extension step. Reaction reagents typically include a thermostable DNA polymerase, deoxyribonucleoside triphosphates (dNTPs), user selected primers for targeting specific sequences, magnesium chloride, and template or target DNA. The process is rapid, requiring between minutes and hours to generate enough discrete sized target sequences for detection; a single thermal cycle may require as little as a few seconds to complete. The length of time required for a reaction is typically a function of variables such as the length of the target sequence and the heating and cooling rates of the thermal cycler used. However, it is now possible to find PCR systems capable of thermal cycling speeds so fast that decreasing cycle time further would not be worthwhile without first finding a DNA polymerase capable of working faster than those currently

4

Molecular Detection of Foodborne Pathogens

in use.13 While it is possible for PCR to routinely detect low copy numbers in a reaction, many reactions use between 5 and 10 µl sample volumes, yielding a lower detection limit of close to 103 CFU/ml.14 Among the expanding array of nucleic acid amplification techniques, PCR remains the most popular method, presumably as a result of its cost and ease of use,15 and has been used extensively for the detection of foodborne pathogens. By the early 1990s numerous primer sets had been developed for the detection of pathogens and the food industry had gained interest in this powerful method.16 The technique was initially reported in 1985,17 explained in full detail in 1986,18 and has since undergone several significant modifications including the use of a thermostable polymerase19 preventing enzyme destruction at denaturation temperatures, and “hot start” enzymes20 for temperature induced activation control, reducing the possibility of nonspecific product formation. Other major advances have included amplicon formation monitoring without opening the reaction tube,21,22 yielding facile quantification of initial target copy numbers, and the use of melting curve analysis to evaluate product specificity, which in some cases, allows extension of the quantifiable range beyond what is possible with threshold cycle analysis alone.23 In addition to evaluating a reaction’s specificity and detection limit, PCR reaction efficiency is often used to assess performance. The number of target copies generated after n cycles, xn, is a function of the initial target copy number xo and the amplification efficiency ε:

xn  = xo(1 + ε)n,

(1.1)

with the amplification efficiency ranging from 0 to 1. Assay parameters that may influence reaction efficiency include primers, annealing temperature, and type of polymerase used. Annealing temperature optimization may be used to balance reaction efficiency and specificity. Approaches to the quantification of real-time PCR products have been described,22,24,25 and techniques typically involve the monitoring of fluorescence accumulation as a function of cycle number through specific or nonspecific dsDNA binding dyes. The threshold cycle, CT, is the fractional cycle at which enough fluorescence has accumulated to rise above the background signal and may be used for quantification. Absolute quantification is possible with unknown samples by running reactions of known template copy numbers to obtain a relationship between the threshold cycle number and the amount of initial template in the reaction. A mathematical model for relative quantification purposes has also been described.26 1.2.2.1.1 Practical Considerations for PCR-based Detection The strength of PCR is its weakness; the assay is incredibly sensitive to the detection of nucleic acids. Since PCR products serve as substrates for subsequent reactions, extremely large numbers of target copies may be generated. As a result

care must be taken in reaction setup and amplicon handling following a reaction to prevent carry-over contamination. Kwok and Higuchi27 list important steps to avoid the occurrence of false positive results, including physically separating the preparation of PCR reagents and the handling of PCR products, as well as frequently changing disposable gloves. While technique is paramount in the ability to generate reproducible results, a brilliant enzymatic approach has also been used to avoid false positive results. Longo et al.28 used a strategy that involved using dUTP in place of dTTP for PCR. All subsequent reactions were treated with uracil DNA glycosylase (UDG), followed by thermal inactivation of this enzyme prior to starting thermal cycling. As a result, any carry-over contaminating DNA would contain uracil and ultimately be rendered unamplifiable through the action of UDG, while simultaneously leaving target DNA intact. Nonspecific amplification arises from primers that bind to unintended targets such as themselves (primer dimers) or other unintended sequences present in the reaction mixture (e.g., DNA sequences from the natural microbiota present in foods). Methods to minimize nonspecific amplification include proper primer design, optimization of assay conditions, and the use of a hotstart DNA polymerase. Wittwer et al.29 studied the influence of annealing time on product specificity. Tests indicated that as annealing time increased, so did the tendency of primer sets to form nonspecific products. Although specificity was generally improved with short annealing times, in some cases there was a tradeoff in the amount of product formed and the specificity of the products formed. Other techniques for optimizing PCR conditions include varying the concentrations of primers and MgCl2, and evaluating two and three step thermal cycling formats. The ultimate test of a primer set’s specificity is in evaluating the performance with target and nontarget DNA sequences. Although it is possible to generate millions of amplicon copies in an hour or less, one complicating factor with PCR testing of food samples is that the level of inhibition is a function of the type of food tested.30 PCR inhibitors may hamper cell lysis, making it difficult to extract DNA, degrade or sequester nucleic acids, or they may act on DNA polymerase.6 In an effort to increase the likelihood of detection when pathogens are present in a sample, separation methods, enrichment procedures, and the extraction of DNA have been used. A negative PCR result could indicate that the target sequence was not present in the reaction or that the reaction itself failed. In order to avoid the uncertainty of such a result in diagnostic PCR, it has been proposed that PCRs contain an internal amplification control.31 Internal amplification controls are nontarget DNA sequences that will be amplified regardless of whether or not the target sequence was present in the reaction. If the internal amplification control is not amplified, then the reaction failed, and it is not possible to know if the target sequence was present in the failed reaction, so the detection step must be repeated.

5

Molecular Detection: Principles and Methods

The amplification of nucleic acids for detection purposes is usually just one step of a procedure that involves assay design and sample preparation prior to amplification, followed by specificity and sensitivity analysis. Some steps prior to and after amplification are shown in Figure 1.1.

using a technique such as agarose gel electrophoresis to ­compare the mobility of standard DNA ladder to the mobility of the amplified DNA (Figure 1.2). Comparison of the known standards to the PCR products can be used to estimate the size of the products formed. This step may be followed by performing a Southern blot to evaluate sequence specificity. Two potential drawbacks to traditional PCR are that (i) endpoint quantification is challenging and (ii) it is necessary to open the reaction tube to verify reaction product specificity,

1.2.2.1.2  Traditional PCR Traditional PCR techniques involve amplification of a target sequence of interest followed by product size verification

Assay design

Sample preparation

• target selection • software aided oligo design

• enrichment • extraction • separation

Specificity analysis

Target amplification • thermal cycling • isothermal incubation

Quantification • CT analysis

• gel electrophoresis • sequence specific probes – Southern blotting – labeled probes – microarray hybridization • complementary sequence Tm • sequencing

Figure 1.1  Steps used for the detection of nucleic acids by amplification. Quantification is not essential to verify specificity.

Melt, anneal primers

Repeat multiple cycles

Extend

Millions of copies of original template

Traditional PCR with gel

Real-time PCR with nonspecific detection

Real-time PCR with sequence-specific detection TaqMan

Molecular Beacon

Unbound probe with reporter and quencher

PCR products

Probe binds Primer binds

PCR proceeds Probe displaced Fluorescent reporter separated from quencher

DNA ladder

Unbound, unactive SYBR green

ssDNA

Bound, active SYBR green

dsDNA

Unbound molecular beacon Target

Probe binds, Beacon unfolds, Reporter activated Reporter, quenched Quencher Reporter, activated

Figure 1.2  Representation of PCR and detection protocols. The principle of PCR is illustrated in the top part of the figure. On the bottom are detection techniques. These include gel electrophoresis after traditional PCR. Real-time PCR with nonspecific fluorescent dye is shown where the dye only fluoresces when associated with double stranded DNA. Real-time PCR with probe-based detection with TaqMan and Molecular Beacon technologies is illustrated where the fluorescent reporter must be physically separated from the quencher in order to fluoresce.

6

increasing the opportunity for carry-over contamination. Traditional PCR therefore requires separate instrumentation for the amplification and evaluation of dsDNA products. However, provided that PCR products are handled carefully and that real-time quantification is not necessary, traditional PCR techniques can be used with great success for the detection of food pathogens. A single enrichment, thermal cycling protocol, set of PCR reagent components and concentrations were used for the detection of 13 foodborne pathogens by Wang et al.32 Agarose gel electrophoresis on 2% agarose gels stained with ethidium bromide was used for separation of PCR products. The PCR detection limits reported ranged from two cells to 5 × 104 cells for E. coli O157:H7 and Shigella spp., respectively. 1.2.2.1.3  Real-time PCR The ability to monitor amplicon accumulation as a reaction proceeds has drastically improved the field of nucleic acid detection. In addition to facilitating the quantification of initial target copy numbers, real-time PCR allows an operator to evaluate product specificity without opening the reaction chamber, saving time, and reducing carry-over contamination risk. Real-time PCR systems offer a wide range of capabilities. These include the ability to handle thousands of samples per day, perform 35 thermal cycles in under 40 minutes, and detect initial target copy numbers over a range from 10 to 1010.33 The design of real-time PCR assays has been aided by commercially available software packages that can determine optimal primer, probe, and reaction conditions, given a specific sequence of interest. Real-time PCR assays are typically designed to target short DNA fragments using primers specifically selected to avoid the formation of primer dimers. The increase in fluorescence in response to amplicon formation is generally accomplished in one of two ways: through the use of a nonspecific dsDNA binding, or by sequence specific probes that generate a signal only in the presence of the target DNA sequence. Realtime PCR techniques and applications have been reviewed extensively,14,34–37 and experimental comparisons among instrumentation and assay formats have been performed to compare sensitivities.38 1.2.2.1.4  Real-time PCR—Nonspecific Detection Nonspecific dsDNA binding dyes have been used for realtime PCR fluorescence based detection systems for target quantification and specificity evaluation. Over the course of thermal cycling, an increase in the amount of fluorescence generated is recorded. The earlier this increase in fluorescence occurs, the larger the initial target copy number present in the reaction. Following thermal cycling, product specificity is verified by slowly raising the reaction temperature through a broad temperature range that includes the expected product melting temperature, while simultaneously recording fluorescence in order to determine the melting temperature (Tm) of any dsDNA products that have formed (Figure 1.2). The melting temperature of a dsDNA

Molecular Detection of Foodborne Pathogens

product is the temperature at which half of the product has become ss. This melting temperature is a function of the dsDNA length, GC content, solution salt concentration, and dye concentration. Advantages of using nonspecific ds binding dyes as compared to probe based systems include cost and ease of assay design. As a result, this approach may be used as a less expensive alternative for initial testing with a primer set. However, this technique does not provide information regarding the length or sequence of the amplified product, and GC rich regions within a single amplicon may create complex melting profiles with multiple peaks.39 SYBR Green is a nonspecific dsDNA binding dye that is frequently used in real-time PCR assays. This dye binds to the minor grove of dsDNA and does not give strong fluorescence when free in solution. SYBR Green real-time PCR assays have been successfully used for the detection of foodborne pathogens, with specificity verification performed by melting curve analysis.40,41 1.2.2.1.5  Real-time PCR—Sequence Specific Detection A large number of real-time PCR strategies that are based on fluorescene increases in response to sequence specific detection have also been developed. Probe based real-time PCR techniques are advantageous over the use of nonspecific dsDNA binding dyes in that they may not require analysis of PCR amplicon melting temperatures for product specificity—fluorescence generation is a function of the probe binding to a specific sequence of DNA. In the case of real-time PCR development with probe based systems, excitation and emission wavelengths of the fluorophores selected must be kept in consideration.34 Sequence specific chemistries that have been incorporated into real-time PCR assays include those based on a sequence specific probe and DNA polymerase exonuclease activity, molecular beacons, and self-quenched hairpin primers. One real-time PCR chemistry (TaqMan®) that has been used extensively for the detection of foodborne pathogens relies upon the 5′ exonuclease activity of Taq polymerase. A probe containing a reporter and quencher in close proximity to one another binds to a target region between the two primers which define the ends of the discrete fragment ultimately formed. This probe is cleaved by the 5′ exonuclease activity of a DNA polymerase, separating the fluorophore and quencher, generating increases in fluorescence as a direct result of specific probe binding and target fragment extension (Figure 1.2). Numerous assays have been developed with this chemistry.42–45 Molecular beacons are stem and loop oligonucleotide structures used for sequence specific detection. The loop portion contains a sequence that is complementary to a chosen target, while the stem portion contains a short sequence of bases at the 3′ and 5′ ends that are complementary to one another but not the target.46 Fluorescence and quenching moieties are attached to the ends of the beacon. The beacons are designed such that with no loop complementary sequence present the stem structure is stable, but in the presence of a

Molecular Detection: Principles and Methods

complementary target sequence the arms of the stem separate. This separation changes the conformation of the beacon to a more stable structure, allowing simultaneous separation of the fluorophore and quencher, leading to fluorescence generation (Figure 1.2).46 Molecular beacons have been used in numerous applications,47 outside of monitoring specific amplicon formation in real-time PCR. Molecular beacons have been used in multiplex PCR applications for the simultaneous detection of four pathogenic retroviruses48 and four V. cholerae genes.49 Hairpin primers have also been used to monitor product formation as a function of cycle number. Blunt end hairpin primers using fluorophores with no quencher molecules were used with great success in a real-time PCR assay.50 Nazarenko et al.50 also demonstrated that these blunt end hairpin primers reduced the formation of primer dimers without PCR template present, thereby showing the outstanding specificity of the system. Nordgren et al.51 used this type of chemistry to detect norovirus (NV) genogroups I and II. Using hairpin primers it was possible to distinguish between genogroups in a duplex PCR through melting curve analysis. 1.2.2.1.6  Reverse Transcriptase PCR Enrichment procedures have successfully been used for the sensitive detection of viable foodborne pathogens, but this technique is time consuming, as it is a function of the target organisms growth. While PCR is capable of detecting low levels of target DNA, DNA detection does not provide information regarding the viability of a cell; food processing may destroy bacteria while leaving behind DNA and this DNA may be present even if its host cell is no longer alive.52 On the other hand, RNA is easily destroyed, which makes it suitable for determining organism viability.30 Reverse transcriptase PCR of mRNA targets has demonstrated that these molecules are indicators of cell viability.53,54 Following RNA purification and degradation of contaminating DNA from a sample of interest, RNA is reverse transcribed and the synthesized complementary DNA or cDNA may be amplified as is typically done for any DNA target. Reverse transcriptase PCR has been used successfully for the detection of foodborne bacterial pathogens55 and viruses.56 A real-time reverse transcription PCR assay using a TaqMan minor grove binding probe was implemented for the quantitative detection of H5 avian influenza down to 100 target copies.57 1.2.2.1.7  Multiplex PCR The amplification of several target sequences in a single reaction tube can be accomplished by optimized multiplex PCR assays. The motivation for such an approach includes cost efficiency58 and a reduction in laboratory effort and time.59 Conditions such as annealing temperature and reagent concentrations must be adjusted to allow for the simultaneous amplification of more than one target. Multiplex PCR optimization may be complicated, resulting in preferential amplification, poor sensitivity, and poor specificity59 if satisfactory conditions for all primer and template combinations cannot

7

be met. In comparison with single PCR reactions, multiplex PCR assay design considerations include designing long primers with higher melting temperatures and using elevated MgCl2 concentrations.60 Additionally, design considerations should include a method to distinguish between amplicons following thermal cycling. Methods may include designing target sequences of different sizes or melting temperatures for discrimination using gel electrophoresis or dissociation analysis with nonspecific dsDNA binding dyes, respectively. Using real-time PCR probes with different excitation and emission wavelengths may also be used to accomplish this goal. Mutliplex PCR has been used to detect multiple gene targets for speciation and virulence determination in Listeria monocytogenes.61 Other multiplex assays have been aimed at detecting food or waterborne pathogens of differing genera.58,62–65 Lee et al.64 simultaneously amplified sequences from Salmonella enterica, Salmonella typhimurium, Vibrio vulnificus, Vibrio cholerae, and Vibrio parahaemolyticus with multiplex PCR from seeded oyster homogenates. Following enrichment and DNA purification, it was possible to detect each pathogen at a level of 102 cells/g of oyster homogenate. Kong et al.63 were able to simultaneously detect Aeromonas hydrophila, Shigella flexneri, Yersinia enterocolitca, Salmonella typhimurium, Vibrio cholerae, and Vibrio parahaemolyticus in marine water with detection limits ranging from 100 to 102 CFU in a total assay time of less than 12 hours. Molecular beacons were used for the simultaneous detection of four retroviral target molecules in the same reaction tube.48 Using different colored fluorophores with emission maxima separated over the visible range and target sequences less than 130 bp, Vet et al.48 detected as few as ten retroviral genomes. 1.2.2.2  Isothermal Amplification Within the last 20 years, many techniques have been developed that allow for amplification of nucleic acids under isothermal conditions. These techniques include loop mediated amplification (LAMP),66 nucleic acid sequence based amplification (NASBA),67 rolling circle amplification (RCA),68 and strand displacement amplification (SDA).69 Isothermal amplification simplifies hardware requirements as compared to PCR in that they do not require a system for thermal cycling, and may even work with a simple water bath setup. These techniques may use several sets of primers or more than one enzyme to carry out amplification of the target product without thermal cycling. NASBA is an isothermal amplification process developed shortly after PCR began gaining widespread attention.67 NASBA is a sensitive detection method for the detection of RNA or DNA. The reaction typically consists of three enzymes including T7 RNA polymerase, deoxyribonucleoside triphosphates, two specific primers, and buffering reagents and takes place at approximately 40°C. One major advantage of the procedure is that contaminating genomic DNA does not create problems with the assay as it will not be

8

amplified due to the fact that there is no thermal denaturation step involved with the process.67 NASBA has been used for the detection of Hepatitis A virus (HAV) using primers targeting major capsid proteins.70 Jean et al. used Northern blotting and dot blot hybridization to verify the specificity of their reaction and found a detection limit of 0.4 ng RNA/ml as compared to a reverse transcriptase PCR assay used that yielded a detection limit of 4 ng RNA/ml. Other NASBA published methods for the detection of pathogens in foods have been listed by Rodríguez-Lázaro et al.71 Real-time NASBA has also been used to show product formation as a function of time. Molecular beacons were used to generate fluorescence signals with NASBA assays for the detection of Vibrio cholerae72 and HAV.73 LAMP is a procedure using four primers that have a total of six binding sites on the target DNA sequence. The isothermal reaction allows for the generation of 109 target sequences in less than one hour.66 A LAMP assay targeting the invA gene of Salmonella was developed by Wang et al.74 using an amplification time of approximately 60 minutes and run at 65°C. The detection limit of the LAMP assay was 100 fg of DNA per reaction, whereas a PCR approach gave a detection limit of 1 pg of DNA per reaction tube. 1.2.2.3  Microarray Detection In addition to Southern blots, gel electrophoresis, melting temperature analysis with nonspecific dsDNA binding dyes, and probe based amplification detection, microarrays have been used to analyze the specificity of PCR products. DNA microarray technology (aka DNA chips or gene chips) involves the placement of user defined oligonucleotide probes in specific locations on a solid substrate such as glass. Following hybridization of target DNA sequences to probes anchored on a chip’s surface, fluorescence detection can be used to monitor binding events. Depending on the sensitivity required, microarrays can be used with or without upstream amplification steps. Software analysis of large data sets that are generated greatly facilitates the process of data analysis. The advantages and limitations of several microarray software packages have been reviewed.75 Microarrays may be an effective way of distinguishing between nonspecific and target product formation and therefore this detection strategy may allow the use of more primers in a multiplex PCR assay than would normally be possible.76 Amplification methods have been used in combination with microarray technology for the detection of E. coli O157:H7.77 Wilson et al.78 were able to specifically detect 18 pathogenic microorganisms including, prokaryotes, eukaryotes, and viruses using PCR in combination with a microarray containing over 50,000 probes and with a detection limit as low as 10 fg of DNA.

1.2.3  Fluorescence in situ Hybridization (FISH) Fluorescence in situ hybridization (FISH) is a technique for the probe-based identification of nucleic acids without

Molecular Detection of Foodborne Pathogens

amplification. The technique can be used to specifically identify microbial cells in environmental samples and rRNA molecules are frequently targeted.79 Fluorescently labeled probes can be used to generate signals in the presence of specific target sequences, seen with fluorescence microscopy. Typical steps include sample preparation by fixation and permeabilisation, probe binding, removal of unhybridized probes by washing, and flow cytometry or microscopy detection.79 A FISH technique for the detection of Listeria monocytogenes showed specific detection of the target microorganism and detection was possible in sheep milk samples.80

1.2.4 Immunological Detection Methods At the core of all immunological assays is an antibody and antigen interaction. Numerous formats have been used to detect these binding events and immunological assays have been widely used for the detection of foodborne pathogens. Assay specificity and sensitivity is a function of the quality and type of antibodies used in binding to specific antigen epitopes. Many immunoassay formats are based on the enzyme linked immunosorbent assay (ELISA).81 ELISAs are commercially available for the detection of foodborne pathogens, and the method can be used for the detection of antibodies or antigens. The technique involves coating an antibody to a solid support surface, adding a sample of interest and incubating, and washing to remove nonspecific interactions. This step is followed by the addition of a second antibody to create a sandwich structure between the primary bound antibody, the target of interest, and this secondary antibody. The secondary antibody may be conjugated with an enzyme or fluorophore for detection and quantification with a plate reader. In this assay format, the target antigen must have at least two antibody binding sites.82 Muhammad-Tahir and Alocilja83 used a sandwich immunoassay with lateral flow disposable membranes and polyaniline-conjugated antibodies, and conductance measurements yielded detection limits of less that 100 CFU/ml. Other methods for evaluating immunological binding events include fluorescence microscopy and surface plasmon resonance (SPR). Fluorescence microscopy has been used to evaluate antibodies against protozoan parasites Giardia and Cryptosporidium.84 SPR sensors measure refractive index changes that result from surface plasmon excitation at the interface between a thin metal film and a dielectric material.85 SPR is attractive because it is a label-free technique, but has sensitivity limitations in terms of the size range of molecules that can be detected. An SPR system was used to detect Salmonella enteritidis and Listeria monocytogenes using antibodies against the pathogens on a gold sensor surface.86 The lower limit of detection was 106 CFU/ml for the pathogens, and it was noted that this sensitivity was comparable to an ELISA using the same antibodies. Immunoassay sensitivity and potential cross reactivity should be carefully considered in comparing detection methods. Another consideration in using immunoassay based systems is that antibodies must be raised against antigens. As a result, immunological methods typically must be used

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Molecular Detection: Principles and Methods

with microorganisms that have been sufficiently characterized.87 The long development times associated with monoclonal antibodies and requirement of in vivo generation makes the widespread application of this technology complicated.88 Also, in some cases it may be difficult to confirm the identity of a microorganism through immunological testing alone. On occasion, reference laboratories found that serotyping could not be used to verify the identity of strains that were initially identified as Salmonella sp.89 Additionally, another nucleic acid based technology may be a suitable alternative for a range of molecular targets traditionally detected by antibodies. Aptamers, single stranded ss DNA or RNA that fold into conformations allowing specific binding to targets, have been proposed as alternative recognition molecules to antibodies.88

1.2.5 Combined Detection Methods Due to limitations of individual detection methods, the combination of two or more techniques has been used for verification purposes, ensuring adequate specificity and sensitivity of results. In a study examining 244 stool samples from an outbreak of gastroenteritis, transmission electron microscopy (TEM), PCR, and ELISA formats were used for the detection of NV.90 The results indicated that at least two of the methods should be used in order to increase the level of confidence in the diagnosis. Combining methods has also been used to enhance the performance of individual assays. Immuno PCR (IPCR) was introduced in 199291 and is a method that can dramatically increase the sensitivity of immunoassays such as the commonly used ELISA. IPCR involves the use of an antibody-DNA conjugate to bind specifically to a target antigen. The antibody is bound to DNA that can then be amplified by PCR. The system is designed such that the presence of PCR product in a reaction means that the target antigen has been detected. One advantage of this technique over other types of PCR methods is that the sequence of DNA to be amplified can be entirely selected by the user.92 An overview of IPCR applications, including pathogen protein detection assays, along with detection limits and sensitivity increases compared to ELISA results, is given by Niemeyer et al.92 A real-time IPCR assay to detect NV capsid proteins in food and fecal samples was developed by Tian and Mandrell.93 They found that PCR inhibitors had a minimal impact on the antigen capture and were removed by wash steps. The real-time IPCR system was the first report to detect NV in contaminated foods without virus purification or concentration. Using a tri-antibody system, the results showed a greater than 1000 fold improvement in sensitivity in comparison to an ELISA assay alone.

1.2.6  Foodborne Pathogen Typing Molecular typing can be used to determine variability within a population of closely related microorganisms and has been

valuable in epidemiological investigations. It is especially important when distinguishing between multiple isolates of the same species. Frequently used methods for studying molecular genetics of bacterial pathogens include pulsedfield gel electrophoresis (PFGE), PCR, and PCR-RFLP (restriction fragment length polymorphism).94 Schwartz and Cantor95 developed PFGE for the separation of large DNA fragments on a 1.5% agarose gel. By alternating the direction of the electrical field across a gel in a perpendicular fashion and varying the pulse length of the different field orientations in a nonuniform fashion from 1 to 90 seconds, it was possible to separate fragments as large as 2000 kb. By changing the direction of the electric field across a gel over short time intervals, it was possible to separate much larger fragments of DNA than was originally possible with standard gel electrophoresis. Whole bacterial chromosomes may be cut by rare digestion enzymes, generating a moderate number of DNA fragments suitable for gel analysis, essentially creating a genetic fingerprint of banding patterns for comparison between strains of the same species.94 PFGE is a technique often used for typing of many bacterial foodborne pathogens and the technique has applicability in studying strain population variability. A typing scheme was created by Wong et al.96 using over 500 strains of Vibrio parahaemolyticus collected from 15 countries and 115 PFGE patterns were identified. It was also found that the restriction enzyme SfiI resulted in clearly separated bands, as opposed to the use of other restriction enzymes.

1.2.7 Microfabrication and Microfluidics Advances in microfluidics along with development of integrated lab-on-a-chip or micro total analysis systems (µTAS) have generated platforms capable of small scale sample preparation, fluid transport, and biological detection.97 Advantages of these microsystems over amplifications on larger scales are that reduced reagent volumes are required, and it may be possible to reduce the amount of time required for the reaction to take place.98 Disadvantages of some microsystems include increased nonspecific binding and the reduction of signal intensity.98 Microchip PCR systems offer advantages of low power consumption as well as rapid heating and cooling. Belgrader et al.99 developed the Advanced Nucleic Acid Analyzer using ten silicon reaction chambers, and detection limit ranges between 102 and 104 organisms/ml were achieved. Neuzil et al.100 obtained heating and cooling rates in excess of 100°C/s using a 100 nl PCR volume with a silicon micromachined chip in a system was able to complete 40 cycles in 5 minutes and 40 seconds.

1.2.8 Other Molecular Detection Approaches Manipulation of nanomaterial properties for targeting biomolecules has created the potential for new techniques that are competitive with ELISA and PCR methods.101 The applications of nanostructures in biodiagnostics has been reviewed.101

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Many silver and gold nanoparticle based ­methods have been used to detect DNA. A label-free platform using silver nanoparticles and smooth silver films was used for the detection of ssDNA by surface-enhanced Raman scattering (SERS).102 In another system, gold nanoparticles were functionalized with thiolated oligonucleotides to detect DNA hybridization by transmission SPR spectroscopy.103 Aptamers are ss nucleic acids that can be generated by a process known as systematic evolution of ligands by exponential enrichment (SELEX), by using libraries of synthetic nucleic acids.88 After folding into a particular conformation, the resulting nucleic acid ligands are capable of specifically binding to a wide range of targets including proteins, making these molecules a potential alternative to antibody based detection. Aptamers are beginning to emerge as molecules that can contend with antibodies in the fields of diagnostics and therapeutics.104 Specific advantages of aptamers over antibodies include their ability to reform their structure following denaturation, an in vitro as opposed to animal or cell based selection process, and chemical synthesis of the selected sequence, making it possible to produce the selected ligand in a very repeatable fashion.104,105 Electrochemical nucleic acid detection techniques have emerged that are label-free and therefore do not require fluorescent dyes and optical components. A disposable electrode system has been used for sensing fM quantities of specific ssDNA sequences, and it was possible to verify hybridization specificity down to a single base pair mismatch by using melting curves.106 Other electrochemical DNA systems have shown promising results in detecting DNA in blood serum107 and PCR products amplified from the gyrB gene of Salmonella typhimurium.108

1.2.9 Assay Design and Data Analysis Software Nucleic acid based detection techniques have grown at a staggering rate due to the availability of target sequence data, powerful methods for nucleic acid amplification, and the ability to easily design suitable nucleic acid sequences for a particular assay. The design of a sensitive, specific PCR assay includes many considerations, and some of the most important are selecting appropriate primers and target DNA sequences. Computer aided PCR assay design systems began appearing not long after the amplification technique was introduced. One such program provided the ability to evaluate DNA duplex stability, oligonucleotide specificity, and oligonucleotide self-complementarity.109 Significant empirical optimization with poorly designed primers can be costly, time consuming, and may not yield adequate results. PCR assay design software can aid in finding primers that have minimal tendency to form secondary structures, closely match primer melting temperatures, find a suitable amplicon size, and predict its melting temperature, all in less time than it takes to select parameter constraints. Additionally, user defined criteria allow primer sets to be rated in terms of their ability to match desired characteristics. Many packages can

Molecular Detection of Foodborne Pathogens

be found online at no charge by using keywords phrases such as “PCR design software” and range in available features from displaying oligonucleotide secondary structure formation to design aides for multiplex real-time PCR assays.

1.3 Detection Targets Just as important as the selection of a suitable method for detection is the selection of an appropriate target to detect. Targets for detection must be unique to the organism of interest. The ideal target would be a gene or a noncoding region with a unique sequence present only in the organism of interest. While unique genes exist, most detection systems take advantage of sequence variations in genes that are shared by many different organisms. Only by studying these organisms, comparing sequence data, and determining specificity have researchers elucidated targets suitable for detection systems in foods. While specificity is an issue, the target should not be so specific that it fails to detect most strains of a species. The detection sequence should be relatively stable within the species. Genes that undergo high rates of recombination, such as some surface antigens that change often to evade immune systems, are not desirable targets. The most common detection sequences are in genetic regions that share some common traits. These loci are common to most if not all the isolates of a species, and they have a high level of sequence conservation, but enough variability in sequence and/or length to distinguish them from similar loci in other genera and sometimes species within genera. Good candidates are genetic loci that are somewhat constrained in sequence because the gene products encode products of essential function but still display some amount of variability (e.g., ribosomal RNA or cytoskeletal proteins). Technical considerations play a role in the choice of detection targets as well. Some high G + C regions may not have high PCR efficiency. This is something to keep in mind if universal primers are being used in a food with a large amount of natural microflora (such as produce or raw meat) when testing for a pathogen that may be present in low numbers. If the template for detection is present in very low numbers, it could be missed if PCR amplifies competing targets with higher efficiency. This is why targets should be tested in laboratory situations with food samples contaminated with pathogens. This allows for assessment not only of the target but the potential inhibition of PCR by components of the food matrix.

1.3.1  Viral Targets The limited genetic information in viruses in relation to the rest of the organisms discussed in this book necessitates a separate discussion of viral detection targets. Viruses consist of genetic material within a proteinaceous capsid and sometimes a surrounding lipoglycoprotein envelope. They are completely dependent on host cells for the expression of their genetic material, their reproduction, and their assembly. Since they have neither organelle structures nor ribosomes,

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Molecular Detection: Principles and Methods

protocols using those targets for detection are useless for viruses. They have very small genomes in comparison with other classes of foodborne pathogens, meaning that there is not a lot of variety in genes to choose for detection purposes. While some concepts of viral detection are shared by other pathogens, some detection targets are unique to viruses. 1.3.1.1 RNA Targets Some of the viruses involved in foodborne outbreaks carry their genetic information as RNA, so reverse transcriptasePCR is used for detection of them. Some RNA viruses contain a gene for an RNA dependent RNA polymerase, which is used for duplication of the viral genetic information. The mutation rate in RNA viruses is much higher than in DNA viruses because of the lack of proofreading ability in RNA dependent RNA polymerase.110,111 This makes it difficult to find stable regions of the genome to use as sequence markers. The polymerase gene itself is one of the few regions of the RNA genome that is relatively conserved. It is an example of a gene that serves an essential function to the virus, so broad changes in sequence that may affect its function are not tolerated. It is used as a detection target for some RNA viruses including hepatitis A, Norwalk virus, and others.112–115 DNA viruses lack such an accommodating gene to use for detection. 1.3.1.2  Viral Structural Genes Capsid proteins are the viral components on display to the environment. They are the major antigenic determinants in viruses, and as such are unique to each virus. These sequence differences among structural proteins present in all viruses is probably the most exploited target for viral detection.113 Capsid genes are used for detection of both DNA and RNA viruses. The V2 and V3 capsid genes in hepatitis A116 have been used for detection of the virus in spiked food samples.117,118 Similarly, primers to rotavirus conserved genome region 9, which contains genes for capsid structure have also been used as targets for detection.118 Strain variability among the different strains of the same virus group results in divergence of capsid gene sequences, and in some viruses the capsid gene sequences are highly mutable. As a result some isolates are not detected by some capsid-directed primer targets. This has been reported for different varieties of Norwalk virus and others.119 Therefore, capsid-designed primers work for detection as long as they target conserved regions in the capsid sequence.120,121 1.3.1.3 Other Viral Targets Noncoding regions in viral genomes, as long as they are conserved and therefore usable between different isolates, have also been used as detection targets for several viruses.122–124 This is the case with the 5′ noncoding region of HAV.125 In other cases unique genes, such as hemagglutinin in avian influenza virus,126 are detection targets. Additionally, since viruses have small genomes multiple isolates of the

same virus type can be sequenced to find unique regions shared among them. Whether or not they are coding regions, these unique sequences can then be used for detection targets for that virus. This procedure was used to find detection targets for Astroviruses.127

1.3.2 Nonviral Targets 1.3.2.1 Ribosomal RNA Genes The most common target for molecular detection is the DNA encoding ribosomal RNA (rRNA). All organisms except for viruses contain these loci. These genetic loci are uniquely suited for diagnostic purposes because they have regions that are very highly conserved in sequence, as well as regions that are divergent. Depending on which regions of the rRNA are targeted they can give different levels of identification from kingdom through genus and species, as well as sometimes differentiating strains within a species.128,129 These regions are also desirable for identification purposes because they share similar physical chromosomal structures (Figure 1.3). Ribosomal RNA in both prokaryotes and eukaryotes is synthesized as one precursor molecule which is then processed to make ribosomes. In prokaryotes the 16S, 23S, and 5S rRNAs are transcribed as one unit also containing a tRNA. In eukaryotes the 18S, 5.8S, and 28S rRNAs are also transcribed as a single unit. In both cases mature rRNAs are made by processing of the primary transcript. Because ribosomes perform an exact function in all living cells the sequence diversity among functional areas of rRNA is highly constrained, but some variation is tolerable. On the other hand, the nonfunctional regions of rRNA loci are under minimal selective pressure, and their sequences and lengths can vary greatly. These differences in rRNA sequence have been used to determine evolutionary relationships between organisms.128 Another benefit of rRNA loci is that they are often present in multiple copies since many ribosomes are necessary for the functioning of growing cells. This means multiple copies of the template sequence for amplification. Beyond the sequence variability in the nonfunctional regions of the rRNA, variable regions are contained within each of the rRNA subunits that provide targets for detection

16S

4S

23S

5S

16S–23S spacer region

28S

5.8S

18S (SSU) ITS

ITS

Figure 1.3  Basic physical map of ribosomal DNA loci in prokaryotes (top) and eukaryotes (bottom). ITS, internal transcribed spacer region, SSU, small subunit RNA.

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of many foodborne pathogens. The 16S, 23S, 18S and 28S subunits have been utilized for detection in most cases. The 16S and 18S RNAs, sometimes referred to as the small subunit RNAs (SSU RNA) in protozoa, have more sequence diversity than the larger subunits (23S and 28S), and as a result the smaller RNAs are used more often for detection.130,131 SSU RNA is a popular target among the protozoa and helminths.132–134 The 16S rRNA sequences have defined bacterial phylogenetic relationships;128 however, for differentiation between species in the same genus the 16S rRNA region is often not discriminating enough because of its low rate of mutation. Even greater diversity in sequence can be obtained by using the spacer regions between the structural subunits. These regions get transcribed as part of the preribosomal RNA, but are cut away later. Since they are not functional RNAs, the spacer regions are not under selective pressure to retain their sequence, but closely related species share similarities in these regions. Because these spacer regions are bound on either side by conserved regions (Figure 1.3), universal primers exist that will bind to the conserved regions and allow their amplification. Specific probes are then used to detect pathogens. Among prokaryotes this spacer region is called the 16S–23S intergenic spacer region (ISR). In eukaryotes, the analogous region is called the internal transcribed spacer region (ITS). ITSs are present in high copy number, and display phylogenetic divergence such that they can show species differentiation.135 The 16S–23S spacer region is widely used to probe for foodborne pathogenic bacteria,131,136 and ITSs are used quite often for detection of fungi and for some helminths.135,137–139 While very useful, sometimes rRNA is not a preferred target. Since all prokaryotes and eukaryotes contain rRNA, primers will amplify regions from many different organisms. If a pathogen is present in low number among normal microflora in foods, then the pathogen target must compete with other templates present in a sample. If the target of interest has a lower PCR efficiency than others present, then the organism of interest may be missed. rRNA has met with mixed results in amplifications from Giardia, for example, because of a high G + C ratio in its 18S rRNA sequence.140 Also, for differentiation of a pathogenic species from nonpathogenic relatives in the same genus, rRNA may not be discriminating enough. 1.3.2.2  Cytoskeleton Proteins Similar to ribosomal DNA, gene sequences for cytoskeletal proteins have been conserved in eukaryotes. These proteins control vital functions such as growth and division of cells, motility, endocytosis, exocytosis, and maintenance of the cell structure. Because these genes arose early in eukaryotic evolution and their sequences have a slow rate of change, they are useful for phylogenetic comparison of species.141,142 These same traits make them useful detection targets, especially if amplification of ITS regions is problematic. Actin and β-tubulin have been used as targets to distinguish between different species in genres of various fungi.143 Giardin, a

Molecular Detection of Foodborne Pathogens

protein associated with the cytoskeleton in Giardia has been used as a detection target.144,145 These proteins, as such, do not exist in prokaryotes, so cytoskeletal targets are limited to detection of eukaryotic pathogens. 1.3.2.3  Virulence and Toxin Genes Among bacteria and fungi there are many cases where a genus consists of pathogenic and nonpathogenic species. Examples are Listeria, Aeromonas, Aspergillus, and Penicillium, to name a few. Often rRNA is not suitable to distinguish between the pathogen and their closely related nonpathogen in the same genus. One way to distinguish them is by assaying for virulence or toxin genes, which are unique to the pathogen genomes. In bacteria, virulence genes are often grouped together on the genome at discrete loci called pathogenicity islands. The altered G + C content of the DNA in many of the pathogenicity islands in relation to the rest of the genome, and repeated sequences on their edges hint that they arrived in these organisms by horizontal transfer.146 Often the pathogens are genetically similar to their sister species except for the pathogenicity islands and other virulence genes. In order to ensure detection of the pathogen in the food sample, and not the innocuous species in the genus, it makes sense to screen directly for the virulence genes or toxins. Virulence gene sequences can be used as specific targets if the gene is unique enough, and the sequence does not vary much between different isolates of a pathogenic species. Hemolysins are a popular target, and they have been used for detection in foods of Shigella, Vibrio, Listeria, Yersinia, Aeromonas, and others.55,147–154 In addition to virulence genes which are involved in the infection process, some organisms produce toxins which are released from the cells. Sometimes these toxins are released as a part of the disease process; however, many organisms release toxins while growing in a food product. Food poisoning is actually caused by reactions to toxins present in food that are made by organisms that grew there. Toxin genes are usually of unique sequence, and so they are used as detection targets. Examples of using toxin genes as detection targets in foods are cereulide, the emetic toxin of Bacillus cereus in rice, botulinum toxin made by Clostridium botulinum in meat and canned corn, enterotoxin made by Staphylococcus aureus in dairy products, and an array of mycotoxins made by fungi such as Alternaria, Aspergillus, Pennicillium, and Fusarium in apples and grains.137,155–162 Other organisms make toxins as part of the disease process once the organism has already grown in the individual, and these types of toxins are also used as detection targets in foods. Examples of these toxins are cytolethal distending toxin in Campylobacter sp. in poultry, and the shiga toxins in Shigella some E. coli strains in meat and dairy products.163–165 1.3.2.4 Unique Genes and Sequences The best detection targets are genes that are absolutely unique to the organism of interest. Failing that, a gene that has unique sequences is desirable. In this section are several examples of unique gene sequences that are neither

Molecular Detection: Principles and Methods

rRNA nor related to virulence and toxicity, but have been found by studies of the physiology of the organisms in question. In Staphylococcus aureus, the nuc gene is a thermostable nuclease.166 While it is not unique to S. aureus, it has sequences in it that will distinguish it from other similar genes. Therefore, it has been used as a detection target for S. aureus.167 The per gene, which encodes perosamine synthetase, has a sequence that is highly conserved among Brucella species, and primers were designed to take advantage of that specificity for detection.168 A unique region in an open reading frame encoding part of the Type III secretion system was utilized as a target to differentiate Burkholderia pseudomallei strains from other bacteria as well as other Burkholderia spp.169 The genus Pseudomonas encompasses a large number of species, some of which are very closely related, so rRNA can be problematic in distinguishing the pathogens from the nonpathogens. The carA gene which encodes carbamoyl phosphate synthase in Pseudomonas sp. was used to distinguish between different species in the genus in meats.170 In order to differentiate between different strains of E. coli sequences in gadA and gadB, which encode glutamate decarboxylase, have been used in artificially contaminated wheat grain.171,172 For the detection of Salmonella in poultry houses the iroB gene, which is absent in the closely related E. coli, was used.173,174 The cpn60 gene (also known as groEL or hsp60), which encodes a heat shock protein in bacteria, contains within it a fragment that has been useful for determining phylogenetic relationships among bacteria. A database of sequences exists to identify organisms found by using this gene as a detection target.175 The rpsU- dnaG- rpoD region is another locus has been used to differentiate between different bacteria. This region encodes proteins involved in the initiations of protein, DNA, and RNA synthesis, and is another example of a locus that has regions that are highly conserved and others that are variable. It has been found to vary between bacteria genera, but to be relatively conserved between species within a genus.176,177 This region was used to distinguish the foodborne pathogen Enterobacter sakazakii from other Enterobacter sp. in infant formula.178 The Toxoplasma B1 gene, which is highly repetitive (35 copies) and highly conserved among various Toxoplasma spp. has been used as a detection target.179,180 The cytoskeletal protein giardin is a major antigenic determinant that is unique Giardia.181 It is also conserved between the different species of Giardia, and as such is a useful detection target. Among the helminths complete genome sequences are not available for many of the genera. However, sequences have been identified by researchers that are unique to some, and these have been used as detection probes. Organisms that have been detected in this fashion include Clonorchis, Opisthorchis, Paragonimus, Taenia, and Fasciola.182–186 1.3.2.5  Insertion Elements In many cases rRNA gene is a fine target for differentiating between genera. However, it is sometimes not discriminating

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enough for the differentiation of species or subspecies. In the case of Mycobacterium avium subsp. paratuberculosis,187,188 IS900 used. This insertion sequence in the genomes of M. avium subsp. paratuberculosis strains allows discretion down to the subspecies level between different M. avium subspecies.189 If simple detection of genus and species is required for the Mycobacterium genus then 16S rRNA is used. Another insertion sequence, IS711 is used as a species specific detection target for Brucella abortus.190 Primers specific for IS407A have been used to differentiate Burkholderia mallei from B. pseudomallei. 1.3.2.6  Mitochondrial Genes In cases where a whole genome sequence is not available for a eukaryotic organism, mitochondrial genome sequences often are available. These organelles, which are present in almost all eukaryotes, contain genomes on the order of 12–20 kb in length that can provide useful targets for detection. For the most part mitochondrial genomes contain the same complement of genes including those coding for proteins needed for oxidative phosphorylation, rRNA, tRNA, as well as noncoding spacer regions. The mitochondrial genome replicates on its own separate from the nucleus, and the coding regions differ at the rate at which they acquire mutations. The noncoding regions have the most variable sequences. Comparisons of mitochondrial genomes have been used to determine phylogenetic relationships between organisms.191,192 Mitochondrial genes are used often for the molecular detection of helminths and some fungi in foods. The mitochondrial gene COX2, which encodes one of the subunits of cytochrome oxidaise, was used as a target for detection of Saccharomyces cerevisiae.193 Cytochrome oxidase and NADH dehydrogenase genes from the mitochondrial DNA is also a popular target for the detection of helminths such as Clonorchis, Opisthorchis, Fasciola, and Diphyllobothrium.194–196 Mitochondrial sequences can be used to differentiate between closely related species in a genus based on size differences in noncoding regions.195 1.3.2.7 Genes for Surface Expressed Markers Similar to the genes that encode capsid proteins in viruses, genes that encode surface markers in other organisms have been used as detection targets. These surface markers are often antigenic determinants, meaning that their coding regions are sufficiently unique for use in detection. However, a problem with surface markers lies in strain variation. In organisms that change surface markers to evade the immune system, detection based on those markers is not useful. Yet bacterial capsule genes are sometimes used to detect Streptococcus species.197 Streptococcus suis contains an extracellular protein factor encoded by the epf gene that is a specific marker for these strains.198 Among Gram-negative bacteria genes encoding lipopolysaccharide markers were used for detection of E. coli, Salmonella, and Vibrio.199 Flagellar genes in bacteria also fall into this category. Many of the flagellar components are expressed on the surface of the cell. They demonstrate unique signatures, so they are also antigenic. In

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some instances flagella serve as virulence factors for bacteria when they are involved in attachment to animal host cells. Several bacterial detection target strategies utilize flagellar genes to detect and differentiate species of Campylobacter,200 E. coli.201 The eap gene in Staphylococcus aureus encodes a cell-surface associated protein that allows adherence of the bacterium to host cells. This gene was described as a potential detection target.202 In Brucella the BCSP-31 gene encodes a cell surface antigen specific to Brucella spp. BCSP-31, and other outer membrane protein encoding genes have been used as Brucella-specific detection targets.203,204 The gene for the oncosphere-­specific protein tso31 has been used as a detection target for the helminth Taenia.205 The oocyst wall protein gene CpR1 from Cryptosporidium is also a target for food detection systems.134,206,207

1.3.3 Using Multiple Targets The best way to screen for a particular pathogen in foods is to use multiple targets for detection. If a combination of rRNA, virulence, and other relevant target genes are used, then there can be a fair amount of confidence that most or all strains of any pathogen can be detected. The most efficient ways of detecting these multiple targets at the same time would be either with a multiplex procedure or with microarray analysis.60,76,199 Panicker et al.149 used both microarrays and multiplex PCR to detect Vibrio spp. in shellfish. Beyond multiple targets and methods, the chances of detection are enhanced with a pre-enrichment step before molecular analysis. This can take the form of a microbiological enrichment, a capture and concentration with immunomagnetic beads that bind to specific pathogens, or both. Just as multiple targets are used for detection of one pathogen, they are also used to detect several pathogens in one test. Screening for several potential pathogens with one protocol saves both time and money. E. coli, L. monocytogenes, and Salmonella typhimurium were each detected in contaminated wheat in one protocol utilizing a microbiological enrichment step followed by multiplex PCR using primers specific to all three bacteria.171 Multiplex PCR was also used for simultaneous detections of Salmonella and Vibrio in shellfish,62 and Yersinia, Staphylococcus, and Shigella in lettuce.65 A microarray was used to detect the mycotoxin biosynthetic genes of Fusarium, Penicillium, and Aspergillus.137 Wilson et al.78 developed a microarray to detect eighteen prokaryotic, eukaryotic, and viral pathogens at once including Brucella, Clostridium, Staphylococcus, Vibrio, Yersinia, and Fusarium.

1.4  Validation Before widespread use of detection methods, they should be validated to make sure that they will detect most or all isolates of a species. False negatives can lead to disastrous consequences, and false positives can lead to costly, unnecessary recalls by manufacturers. A detection target is not very useful

Molecular Detection of Foodborne Pathogens

if a subset of strains of that organism do not possess it. The best methods have been validated by use in multiple laboratories to make sure they work with different sets of workers. This type of interlaboratory validation is routinely done. For example, 12 laboratories recently validated a Campylobacter detection protocol in spiked chicken carcass rinses,208 and several labs were involved in detection of hepatitis A from spiked food samples.117 Experiments within the food matrix are very important as they will not only assess the detection method within in the presence of potential inhibitors, but also determine how well the target organism competes with natural microbiota in the food. The competition aspect is important to test if an enrichment is involved prior to actual detection. Targets should be screened to make sure they do not react with unwanted organisms. Potential primers and probes should be tested against multiple, closely related species for cross-reactivity. Also during these validations, sensitivity of the method can be determined with known levels of organisms added to foods.

1.5  Conclusions Molecular biology has rapidly revolutionized food diagnostics, driven by biotechnology advances fueled by basic science.209 Numerous molecular detection techniques have emerged that are rapid, sensitive, and specific in detecting nucleic acid sequences of foodborne pathogens. Several factors have led to an explosive growth in methods available for the detection and quantification of foodborne pathogens including the ready availability of synthetic oligonucleotide sequences of approximately 100 bp or less, the ability to modify these sequences at their 3′ or 5′ ends with fluorescence labels or conjugation chemistries, the development of extremely sensitive DNA amplification techniques such as PCR, software tools for molecular assay design and data evaluation, and DNA sequence databases to allow for the search for efficient target sequences. Rapid and simple sample preparation techniques, simplification of data analysis, standardization of molecular testing procedures, and identification of suitable detection targets will lead to the more widespread acceptance of molecular techniques. Additionally, cost effective solutions to pathogen detection and systems for on-site analysis that require minimal operator interface would benefit many industries. Microfabrication, microfluidics, and nanoparticle conjugation chemistries are likely to play significant roles in future systems. The extent to which foodborne pathogen detection solutions will converge upon widely accessible integrated instrumentation solutions that merge preparation, detection, and data interpretation capabilities in a seamless platform remains to be seen. However, molecular detection techniques will likely continue to simplify and increase the speed of detection procedures while simultaneously improving the sensitivity and specificity required for tracking pathogens in environmental, clinical, and food matrices.

Molecular Detection: Principles and Methods

Acknowledgments The authors wish to thank J. Barak, M. Borucki, D. Flaherty, W. Miller and J. Palumbo for helpful discussions and advice.

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Molecular Detection: Principles and Methods 163. Samosornsuk, W. et al. Evaluation of a cytolethal distending toxin (cdt) gene-based species-specific multiplex PCR assay for the identification of Campylobacter strains isolated from poultry in Thailand. Microbiol. Immunol., 51, 909, 2007. 164. Chen, S. et al. An automated fluorescent PCR method for detection of shiga toxin-producing Escherichia coli in foods. Appl. Environ. Microbiol., 64, 4210, 1998. 165. Cebula, T.A., Payne, W.L. and Feng, P. Simultaneous identification of strains of Escherichia coli serotype O157:H7 and their shiga-like toxin type by mismatch amplification mutation assay-multiplex PCR. Appl. Environ. Microbiol., 33, 248, 1995. 166. Alarcón, B., Vicedo, B. and Aznar, R. PCR-based procedures for detection and quantification of Staphylococcus aureus and their application in food. J. Appl. Microbiol., 100, 352, 2006. 167. Barkstad, O.G., Aasbakk, K. and Maeland, J.A. Detection of Staphylococcus aureus by polymerase chain reaction amplification of the nuc gene. J. Clin. Microbiol., 30, 1654, 1992. 168. Bogdanovich, T. et al. Validated 5’ nuclease PCR assay for rapid identification of the genus Brucella. J. Clin. Microbiol., 42, 2261, 2004. 169. Novak, R.T. et al. Development and evaluation of a realtime PCR assay targeting the type III secretion system of Burkholderia pseudomallei. J. Clin. Microbiol., 44, 85, 2006. 170. Ercolini, D. et al. Simultaneous detection of Pseudomonas fragi, P. lundensis, and P. putida from meat by use of a multiplex PCR assay targeting the carA gene. Appl. Environ. Microbiol., 73, 2354, 2007. 171. Kim, J. et al. Simultaneous detection by PCR of Escherichia coli, Listeria monocytogenes, and Salmonella typhimurium in artificially inoculated wheat grain. Int. J. Food Microbiol., 111, 21, 2006. 172. McDaniels, A.E. et al. Confirmational identification of Escherichia coli, a comparison of genotypic and phenotypic assays for glutamate decarboxylase and β-D-glucuronidase. Appl. Environ. Microbiol., 62, 3350, 1996. 173. Bäumler, A.J., Heffron, F. and Reissbrodt, R. Rapid detection of Salmonella enterica with primers specific for iroB. J. Clin. Microbiol., 35, 1224, 1997. 174. Soumet, C. et al. An immunoconcentration-PCR assay to detect Salmonella in the environment of poultry houses. Int. J. Food Microbiol., 48, 221, 1999. 175. Hill, J.E., Town, J.R. and Hemmingsen, S.M. Improved template representation in cpn 60 polymerase chain reaction (PCR) product libraries generated from complex templates by application of a specific mixture of PCR primers. Environ. Microbiol., 8, 741, 2006. 176. Erickson, B.D. et al. Nucleotide sequence of the rpsU-dnaGrpoD operon from Salmonella typhimurium and a comparison of this sequence with the homologous operon of Escherichia coli. Gene, 40, 67, 1985. 177. Versalovic, J. et al. Conservation and evolution of the rpsUdnaG-rpoD macromolecular synthesis operon in bacteria. Mol. Microbiol., 8, 343, 1993. 178. Seo, K.H. and Brackett, R.E. Rapid, specific detection of Enterobacteri sakazakii in infant formular using a real-time PCR assay. J. Food Prot., 68, 59, 2005. 179. Lin, M.-H. et al. Real-time PCR for quantitiative detection of Toxoplasma gondii. J. Clin. Microbiol., 38, 4121, 2000. 180. Burg, J.L. et al. Direct and sensitive detection of a pathogenic protozoan, Toxoplasma gondii, by polymerase chain reaction. J. Clin. Microbiol., 27, 1787, 1989.

19 181. Faubert, G. Immune response to Giardia duodenalis. Clin. Microbiol. Rev., 13, 35, 2000. 182. Wonratanacheewin, S. et al. Development of a PCR-based method for the detection of Opisthorchis veverrini in experimentally infected hamsters. Parasitology, 122, 175, 2001. 183. Parvathi, A. et al. Clonorchis sinensis: development and evaluation of a nested polymerase chain reaction (PCR) assay. Exp. Parasitol., 115, 291, 2007. 184. Kaplan, R.M. et al. A repetitive DNA probe for the sensitive detection of Fasciola hepatica in infected snails. Int. J. Parasitol., 25, 601, 1995. 185. Maleewong, W. et al. Detection of Paragonimus heterotremus in experimentally infected cat feces by antigen captureELISA and by DNA hybridization. J. Parasitol., 83, 1075, 1997. 186. González, L.M. et al. Differential diagnosis of Taenia saginata and Taenia solium infection by PCR. J. Clin. Microbiol., 38, 737, 2000. 187. Pillai, S.R. and Jayarao, B.M. Application of IS900 PCR for detection of Mycobacterium avium subsp. paratuberculosis directly from raw milk. J. Dairy Sci., 85, 1052, 2002. 188. Ravva, S.V. and Stanker, L. H. Real-time quantitative PCR detection of Mycobacterium avium subsp. paratuberculosis and differentiation from other mycobacteria using SYBR green and TaqMan assays. J. Microbiol. Methods, 63, 305, 2005. 189. Bull, T.J. et al. Characterization of IS900 loci in Myco­ba­ cterium avium subsp. paratuberculosis and development of multiplex PCR typing. Microbiology 146, 2185, 2000. 190. O’Leary, S., Sheahan, M. and Sweeney, T. Brucella abortus detection by PCR assay in blood, milk and lymph tissue of serologically positive cows. Res. Vet. Sci., 81, 170, 2006. 191. Boore, J.L. Animal mitochondrial genomes. Nucleic Acid Res., 27, 1767, 1999. 192. Le, T.H., Blair, D. and McManus, D.P. Mitochondrial DNA sequences of human schistosomes: the current status. Int. J. Parasitol., 30, 283, 2000. 193. López, V. et al. A simplified procedure to analyse mitochondrial DNA from industrial yeasts. Int. J. Food Microbiol., 68, 75, 2001. 194. Le, T.H. et al. Clonorchis sinensis and Opisthorchis viverrini: development of a mitochondiral-based multiplex PCR for their identification and discrimination. Exp. Parasitol., 112, 109, 2006. 195. Kim, K.-H. et al. Characterization of the complete mitochondrial genome of Diphyllobothrium nihonkaiense (Diphyllobothriidae: Cestoda) and development of molecular markers for differentiating fish tapeworms. Mol. Cells, 23, 379, 2007. 196. Cucher, M.A. et al. PCR diagnosis of Fasciola hepatica in field-collected Lymnaea columella and Lymnaea viatrix snails. Vet. Parasitol., 137, 74, 2006. 197. Marois, C. et al. Multiplex PCR assay for detection of Streptococcus suis species serotypes 2 and 1/2 in tonsils of live and dead pigs. J. Clin. Microbiol., 42, 3169, 2004. 198. Wisselink, H.J., Joosten, J.J. and Smith, H.E. Multiplex PCR assays for simultaneous detection of six major serotypes and two virulence-associated phenotypes of Streptococcus suis in tonsillar specimens from pigs. J. Clin. Microbiol., 40, 2922, 2002. 199. Morin, N.J., Gong, Z. and Li, X.-F. Reverse transcription-multiplex PCR assay for simultaneous detection of Escherichia coli O157:H7, Vibro cholerae O1, and Salmonella typhi. Clin. Chem., 50, 2037, 2004.

20 200. Harmon, K.M., Ransom, G.M. and Wesley, I.V. Diff­erentiation of Campylobacter jejuni and Campylobacter coli by polymerase chain reaction. Mol. Cell. Probes, 11, 195, 1997. 201. Gannon, V.P.J. et al. Use of the flagellar H7 gene as a target in multiplex PCR assays and improved specificity in identification of enterhemorrhagic Escherichia coli strains. J. Clin. Microbiol., 35, 656, 1997. 202. Hussain, M. et al. eap gene as novel target for specific identification of Staphylococcus aureus. J. Clin. Microbiol., 46, 470, 2008. 203. Imaoka, K. et al. Simultaneous detection of the genus Brucella by combinatorial PCR. Jpn. J. Infect. Dis., 60, 137, 2007. 204. Tantillo, G.M., Di Pinto, A. and Buonavoglia, C. Detection of Brucella spp. in soft cheese by semi-nested polymerase chain reaction. J. Dairy Res., 70, 245, 2003.

Molecular Detection of Foodborne Pathogens 205. Mayta, H. et al. Nested PCR for specific diagnosis of Taenia solium taeniasis. J. Clin. Microbiol., 46, 286, 2008. 206. Laberge, I. et al. Detection of Cryptosporidium parvum in raw milk by PCR and oligonucleotide probe hybridization. Appl. Environ. Microbiol., 62, 3259, 1996. 207. Wiedenmann, A., Krüger, P. and Botzenhart, K. PCR detection of Cryptosporidium parvum in environmental samples--a review of published protocols and current developments. J. Ind. Microbiol. Biotechnol., 21, 150, 1998. 208. Josefsen, M.H. et al. Validation of a PCR-based method for detection of food-borne thermotolerany Campylobacters in a multicenter collaborative trial. Appl. Environ. Microbiol., 70, 4379, 2004. 209. Feng, P. Impact of molecular biology on the detection of foodborne pathogens. Mol. Biotechnol., 7, 267, 1997.

Section I Foodborne Viruses

2 Adenoviruses Charles P. Gerba

University of Arizona

Roberto A. Rodríguez

University of North Carolina

Contents 2.1 Introduction......................................................................................................................................................................... 23 2.1.1 Classification and Morphology................................................................................................................................ 23 2.1.2 Biology, Pathogenesis, and Medical Importance..................................................................................................... 24 2.1.2.1 Gastroenteritis........................................................................................................................................... 25 2.1.2.2 Respiratory Infections............................................................................................................................... 25 2.1.2.3 Pharyngoconjunctival Fever (PCF)........................................................................................................... 25 2.1.2.4 Eye Infections............................................................................................................................................ 25 2.1.2.5 Obesity...................................................................................................................................................... 25 2.1.2.6 Morbidity and Mortality............................................................................................................................ 25 2.1.2.7 Impact on the Immunocompromised........................................................................................................ 25 2.1.2.8 Water- and Foodborne Outbreaks............................................................................................................. 25 2.1.2.9 Occurrence in Water................................................................................................................................. 26 2.1.2.10 Occurrence in Shellfish............................................................................................................................. 26 2.1.2.11 Survival in the Environment and Shellfish............................................................................................... 26 2.1.3 Identification and Diagnosis.................................................................................................................................... 27 2.1.3.1 Culture-Based Techniques........................................................................................................................ 27 2.1.3.2 Antibody-Based Methods.......................................................................................................................... 27 2.1.3.3 Nucleic Acid Probes.................................................................................................................................. 27 2.1.3.4 PCR-Based Techniques............................................................................................................................. 27 2.1.3.5 Integrated Techniques............................................................................................................................... 27 2.2 Methods............................................................................................................................................................................... 28 2.2.1 Sample Preparation.................................................................................................................................................. 28 2.2.2 Detection Procedures............................................................................................................................................... 29 2.3 Conclusions and Future Perspectives.................................................................................................................................. 30 References.................................................................................................................................................................................... 31

2.1  Introduction The potential for the transmission of adenoviruses by foods has only recently received attention and has been almost entirely focused on shellfish. Transmission of adenovirus eye and respiratory infections by recreational waters, however, has been well documented. Recent outbreaks have suggested that adenoviruses can also be transmitted by drinking water. Association of adenoviruses with water and food outbreaks is difficult because of the wide variety of illnesses that the viruses can cause, and of large number of asymptomatic cases. This is exacerbated by the fact that being a nonreportable disease, adenovirus infection is often associated with illnesses not considered foodborne (respiratory infections). Still occurrence of these viruses in food and water should be taken as an indication of their potential to be transmitted by

these routes. The application of molecular methods has been key in our understanding of exposure by food and water.

2.1.1 Classification and Morphology The human adenoviruses belong to the genus Mastadenovirus in the family Adenoviridae and consist of at least 51 serotypes. These serotypes are divided into six subgenera labeled A through F. Each serotype is distinguished by its resistance to neutralization by antisera to other known adenovirus serotypes.1 Table 2.1 outlines the current classification scheme for human adenovirus serotypes. Adenoviruses have a nonenveloped, icosahedral virion that consists of a core containing linear double-stranded DNA (26–45 kb) enclosed by a capsid.2 The capsid is composed of 252 capsomers, 240 of which are hexons and 12 of which 23

24

Molecular Detection of Foodborne Pathogens

are pentons. Each penton projects a single fiber that varies in length for each serotype, an exception being the pentons of the enteric adenoviruses (serotypes 40 and 41) that project two fibers.1 Adenoviruses are approximately 70–100 nm in diameter.

2.1.2 Biology, Pathogenesis, and Medical Importance Due to their physical, chemical, and structural properties, adenoviruses may survive extended periods of time outside host cells. They are stable in the presence of many physical and chemical agents, as well as adverse pH conditions. For example, adenoviruses are resistant to lipid solvents due to the lack of lipids within their structure.4 Infectivity is optimal between pH 6.5 and 7.4; however, the viruses can withstand pH ranges between 5.0 and 9.0. Adenoviruses are heat resistant (particularly type 4) and may remain infectious after freezing.5 Table 2.1 Human Adenovirus Serotype Classification Subegenera A B C D E F

Serotypes 12, 18, 31 3, 7, 11, 14, 16, 21, 34, 35, 50 1, 2, 5, 6 8–10, 13, 15, 17, 19, 20, 22–30, 32, 33, 36–39, 42–49, 51 4 40, 41

Source: Adapted from Shenk, T., Adenoviridae: The viruses and their replication. In Fields Virology, 4th edn. Knipe, D.M. et al. (Eds.), Lippincott Williams and Wilkins, Philadelphia, PA, 2001, and van Regenmortel, M.H.V. et al., Virus Taxonomy: Seventh Report of the International Committee on Taxonomy of Viruses, Academic Press, San Diego, CA, 2000.

Routes of infection include the mouth, nasopharynx and the ocular conjunctiva. Less frequently, the virus can become systemic and affect the bladder, liver, pancreas, myocardium or central nervous system.6  Of the 51 currently recognized human serotypes (a serotype 52 has been proposed), only onethird are associated with a specific human disease (Table 2.2). Other infections remain largely asymptomatic. Adenoviruses are associated with a variety of types of clinical illnesses involving almost every human organ system. Illnesses include upper (pharyngitis and tonsillitis) and lower (bronchitis, bronchiolitis, and pneumonia) respiratory illnesses, conjunctivitis, cystitis, and gastroenteritis. Several studies have found that the enteric adenoviruses are second only to rotaviruses as the causative agents of acute gastroenteritis in infants and young children.7,8 Most illnesses caused by adenoviruses are acute and self-limiting. Although the symptomatic phase may be short, all adenoviruses can remain in the gastrointestinal tract and continue to be excreted for an extended period of time. Species within subgenera C may continue to be excreted for months or even years after disease symptoms have resolved. Adenoviruses can remain latent in the body (in tonsils, lymphocytes and adenoidal tissues) for years and be reactivated under certain conditions, such as a change in immune status. The long-term effect of such a latent infection is unknown.5 Adenovirus infections may be accompanied by diarrhea, though the virus can be excreted even if diarrhea is not present.7 A large proportion of infections caused by subgenera A and D tend to be asymptomatic, whereas the species within subgenera B and E tend to result in a higher rate of symptomatic respiratory illnesses. Immunity is species-specific. The presence of pre-existing antibodies resulting from a previous infection is usually protective. It is difficult to confidently link all adenoviruses to specific illnesses because many infections may be asymptomatic, healthy people can shed viruses.5 Occurrence studies ­comparing infection in healthy and ill people have found

Table 2.2 Common Illnesses Associated with Human Adenoviruses Disease Acute febrile pharyngitis Pharyngoconjunctival fever Acute respiratory disease Pneumonia Epidemic keratoconjunctivitis Follicular conjunctivitis Gastroenteritis/diarrhea Urinary tract Colon Hepatitis

Individual at Risk Infants, young children School-aged children Military recruits Infants, young children, military recruits Any Infants, young children Infants, young children Bone marrow, liver or kidney transplant recipients, AIDS victims or immunosuppressed

Serotypes 1–3, 5–7 3, 7, 14 3, 4, 7, 14, 16, 21 1–3, 4, 6, 7, 14, 16 8–11,13, 15, 17, 19, 20, 22–29, 37 3, 7 18, 31, 40, 41 34,35 42–49 1, 2, 5

Source: Adapted from Horwitz, M.S., Adenoviruses. In Fields Virology, 4th edn. Knipe, D.M. et al. (Eds.), Lippincott, Williams and Wilkins, Philadelphia, PA, 2001; and Enriquez, C.E., Adenoviruses. In Encyclopedia of Environmental Microbiology, Bitton, G. (Ed.), John Wiley and Sons, New York, 2002.

Adenoviruses

25

between 0 and 20% of asymptomatic people can shed adenovirus.

complete recovery. Ad3 and Ad7 are the most commonly isolated species.6

2.1.2.1 Gastroenteritis Estimates of the incidence of adenovirus gastroenteritis in the world have ranged from 1.5 to 12%. Enteric ­adenoviruses are second only to rotaviruses as the leading causes of childhood gastroenteritis.7,9 Diarrhea is usually associated with fever and can last for up to two weeks. Though diarrhea can occur during infection by any type of adenovirus, Ad40 and Ad41 of subgenus F specifically cause gastroenteritis and diarrhea. Adenovirus type 31 (Ad31) is also suspected of causing infantile gastroenteritis. Some estimate that Ad40/41 contribute from 5 to 20% of hospitalizations for diarrhea in developed countries.10

2.1.2.5 Obesity There is accumulating evidence that several viruses may be involved in obesity in animals and humans.14 Studies in chickens, mice and nonhuman primates indicate that Ad36 can cause obesity.15 Obese humans have a higher prevalence of serum antibodies to Ad36 than lean humans.16 Other adenoviruses are capable of causing obesity in animals, but no correlation with antibodies has been demonstrated.16 The metabolic and molecular mechanisms of how adenovirus infections cause obesity are not precisely understood; however, increases in food intake alone cannot explain the observed increases in adiposity (tendency to store fat), suggesting that Ad36 induces metabolic changes.17 One mechanism appears to be that Ad36 influences the differentiation of preadipocyte.17

2.1.2.2 Respiratory Infections Over 5% of respiratory illnesses in children younger than 5 years of age are due to adenovirus infections.11 The initial transmission of adenoviruses is through the nasopharynx. Secondary transmission in households can be as high as 50% due to fecal-oral transmission from children shedding virus in the feces. Adenoviruses can be recovered from the throat or stool of an infected child for up to three weeks.6 Adenovirus respiratory infections are also well documented in adults. 2.1.2.3  Pharyngoconjunctival Fever (PCF) Pharyngoconjunctival fever (PCF) refers to a syndrome of pharyngitis, conjunctivitis, and spiking fever.5 Symptoms of this syndrome include unilateral or bilateral conjunctivitis, mild throat tenderness, and fever. The illness usually lasts from 5 to 7 days, with no permanent eye damage.4 The most commonly isolated adenovirus serotype is 3, although 7 and 14 have also been associated.6 The disease is best known for centering on summer camps, pools, and small lakes.12,13 Transmission of the agent appears to require direct contact with the water, allowing the virus direct contact with the eyes or upper respiratory tract. Secondary spread is common, although adults contracting the disease tend to have milder symptoms, usually only conjunctivitis. 2.1.2.4 Eye Infections Epidemic keratoconjunctivitis (EKC) is a syndrome which causes inflammation of the conjunctiva and cornea. EKC was once referred to as “Shipyard Eye,” as it was first described in shipyard workers.6 EKC is considered highly contagious and begins with edema of the eyelids, pain, shedding tears, and photophobia. Serotypes 8, 11, 19, and 37 can cause EKC. Transmission occurs through direct contact with eye secretions from an infected person as well as through contact with contaminated surfaces, eye instruments, ophthalmic solutions, towels, or hands of medical personnel. Outbreaks have involved mostly adults. Follicular conjunctivitis is often contracted by swimming in inadequately chlorinated swimming pools or in lakes during the summer.6 Most cases result in only mild illness and

2.1.2.6  Morbidity and Mortality Since adenovirus is not a reportable disease agent, there are no national or population-based morbidity and mortality ­figures available; most of the epidemiological data come from the study of select populations who appear to be most affected by adenovirus exposure. These include children in institutions such as hospitals and daycare centers, military recruits, immunocompromised individuals, and groups of families. Enteric infection in children results in disease 50% of the time. This percentage is greater when the infection is centered in the respiratory tract.5 Attack rates for waterborne outbreaks have been as high as 67% in children, with secondary attack rates (person-to-person transmission) of 19% for adults and 63% for children.5 2.1.2.7  Impact on the Immunocompromised Although adenovirus infection may result in mild or asymptomatic infections in the immuno competent in the immunocompromised, the virus can disseminate into any body system and cause pneumonitis, meningoencephalitis, hepatitis (especially in liver and bone marrow transplant patients), and hemorrhagic cystitis (especially in kidney transplant patients).5 According to Hierholzer18 over 11% of transplant recipients become infected with adenoviruses, with an 18% case fatality rate. The enteric adenoviruses are rarely isolated from immunocompromised patients with gastroenteritis or diarrhea and are generally not associated with serious illness in the immunocompromised. 2.1.2.8 Water- and Foodborne Outbreaks Although adenoviruses have been detected in shellfish no foodborne outbreaks have been documented to date. This may be a simple reflection that the virus is never considered a cause and thus no testing is perform to assess if adenoviruses could be involved. However, contact with recreational water has been associated with numerous adenovirus outbreaks over the years. Adenoviruses are the most reported cause of swimming pool outbreaks associated with viruses. Many outbreaks of PCF from nonenteric adenoviruses have come from people

26

swimming in pools and lakes. Ad3, Ad4, Ad7, and Ad14 have been associated with outbreaks in swimming pools19,20 and the adenoviruses have been detected in pool waters after outbreaks.12 It is clear that nondisinfected or inadequately disinfected recreational water is a source of adenovirus infection in swimmers. A routine monitoring of chlorinated swimming pools in South Africa demonstrated the presence of adenovirus by polymerase chain reaction (PCR) in 26 of 93 (15.4%) samples.20 While the detection method did not assess virus viability, it did demonstrate the widespread occurrence of adenoviruses in swimming pools. There have been three drinking water outbreaks reported in Europe in which enteric adenoviruses may have been a cause of gastroenteritis.21–23 Multiple viral agents were involved and the water had not been adequately disinfected. 2.1.2.9 Occurrence in Water Limited data has been available on the occurrences of adenoviruses in water. Only since the development of molecular methods for the direct detection of adenoviruses in water with confirmation tests performed in cell culture has data become available. Adenoviruses have been isolated from wastewater and river water, often more frequently and at higher concentrations than the enteroviruses.24–27 Adenoviruses have also been detected in sewage, rivers, oceans, swimming pools. Adenoviruses are commonly detected in raw and nondisinfected secondary sewage discharges, although little published data is available for the U.S. In Spain, monthly samples of raw sewage, effluent, river water, and seawater were tested using nested PCR amplification. Adenovirus was detected in 14 of 15 sewage, two of three effluent, 15 of 23 river water, and seven of nine seawater samples. Samples that were positive for enterovirus or hepatitis A were also positive for adenovirus, but there was no correlation between the fecal coliform level and adenovirus occurrence.26 In Greece, 36 samples of sewage effluent were tested over a 15-month period using cell culture. Adenovirus was detected in all samples with concentrations ranging from 70 to 3200 cytopathic units (CPU)/l. In Australia,24 raw sewage, primary effluent, and secondary effluent were sampled over a year using cell culture; 25 of 26 raw sewage, 23 of 26 primary effluent and 23 of 26 secondary effluent samples were positive for adenovirus.28 The mean concentrations in sewage, primary effluent, and secondary effluent were 1950, 1350, and 250 infectious units (IU)/l, respectively. Enteroviruses were removed to a greater extent than adenoviruses by activated sludge treatment.24 Both respiratory and enteric adenoviruses have been isolated from surface waters worldwide. Nevertheless, survey data is limited in the U.S. An evaluation of 29 surface water samples in the U.S. yielded 38% positive for infectious Ad40 and Ad41.29 The concentration of Ad40/41 ranged from 1.03 to 3.23 per 100 l. In this study, adenoviruses were more common in surface waters than enteroviruses and astroviruses. Likewise, when comparative studies have been conducted, adenoviruses usually outnumber enteroviruses in surface waters. Infectious adenoviruses have been detected in conventionally treated and disinfected drinking water in Africa and Asia

Molecular Detection of Foodborne Pathogens

using genome detection with PCR in cell culture.30,31 In both of these studies, adenoviruses were commonly detected in the raw, untreated surface water. In one study, adenoviruses were found in 4.4% of the finished drinking water samples that met the current acceptable bacteriological standards. In the other study, adenoviruses were detected at concentrations ranging from 0 to 0.9 most probable number (MPN)/100 l. In the van Heerden et al. study it was noted that none of the adenoviruses growing in cell culture produced cytopathic effects (CPE). 2.1.2.10 Occurrence in Shellfish Interest in the occurrence of adenoviruses in shellfish largely stems from their potential as indicators of other enteric viruses and fecal pollution.26 As with sewage and surface waters adenoviruses appear to be in greater numbers or at least isolated more commonly in shellfish than other enteric viruses, which may be a reflection of their greater stability in the environment.32 In addition, being a DNA virus eliminates the added steps for detection enteric RNA viruses needed for reverse transcription-PCR (RT-PCR). Although cell culture has been used to a limited degree PCR has been the method of choice, because of the long incubation times required for production of cytopathogenic effects in cell culture required for adenoviruses. Adenoviruses have been reported from every continent where shellfish are harvested. In Spain reported that of the mussels and oysters tested 47% contained adenoviruses, 19% enteroviruses, and 24% hepatitis A virus.33 In a multilaboratory study of virus contamination of shellfish in Europe human adenoviruses detected by PCR where found to correlate with the presence of other human enteric viruses and suggested they could be useful as a molecular index of viral contamination of shellfish.34 In a study in Korea adenoviruses was detected in 89% of the oysters collected from several locations.35 In India adenoviruses were detected in 17% of the oysters and 27% of clam samples, but noroviruses and hepatitis A virus were detected.36 However, enteroviruses were isolated with a greater frequency, 37% for oysters and 46% for clams. 2.1.2.11 Survival in the Environment and Shellfish Limited data suggests that adenoviruses survive longer in water than enteroviruses and hepatitis A virus.37 Adenoviruses also exhibit greater thermal stability than enteroviruses. This may explain their longer survival in water. They are capable of surviving for months in water, especially at low temperatures. The double-stranded DNA that comprises the genome of the virus may provide more stability in the environment. In addition, adenoviruses may use host cell repair enzymes to repair damaged DNA. This may also prolong their survival in the environment and enhance their resistance to inactivation by ultraviolet light. Qualitative PCR analyses of adenovirus DNA in oysters and mussels demonstrated that Ad35 could be detected for 6–8 weeks.38 In contrast the virus was detected in cell culture for 4–6 weeks.

Adenoviruses

2.1.3 Identification and Diagnosis 2.1.3.1  Culture-Based Techniques Adenovirus subgenera A through E can be cultured in human cell lines, albeit slowly,39,40 and thus may be overgrown by other faster growing viruses. They also require more than one passage in cell culture for expression of CPE. For this reason usually TCID50 or MPN methods are used for their quantification in environmental samples27 Guanidine can be added to cultures to selectively suppress enteroviruses while allowing adenoviruses to grow.25 A variety of cell lines have been used to grow and/or detect adenovirus such as HeLa cells,24,41 HEp-2 cells,28,41 293 cells,42 Chang conjunctival cells,43 CaCo-2 cells,44 and PLC/PRF/5 cells.43 Hurst et al.25 found that the number of infectious adenoviruses obtained by observing CPE in the 293 cell line was five-fold greater than the number detected via CPE in HEp-2 cells with sewage samples. Based on these findings and those of Takiff et al.45 they suggested that HEp-2 cells might not be as appropriate for detecting Ad40 and Ad41 as 293 cells. The use of HEp-2 cells might miss the enteric adenoviruses that may constitute up to 80% of the adenoviruses found in raw sewage. This might explain the findings of Tani et al.41 who, unlike other researchers, detected adenoviruses at much lower numbers than the enteroviruses in sewage, but relied on the use of the HeLa and HEp-2 cell lines. Grabow et al.43 determined that the PLC/PRF/5 liver cell line was more sensitive for detecting Ad41 and also exhibited CPE earlier than 293 cells and Chang conjunctival cells; however, while Ad40 may be grown using the PLC/PRF/5 cell line, CPE is not observed.43 This cell line has been used to study the survival and recovery, respectively, of Ad40 and Ad41 in water.19,37 also reported that the PLC/PRF/5 cell line was at least as sensitive as the HEp-2 cell line for isolating the lower-numbered serotypes (i.e., Ad1, Ad2, Ad3, Ad5, Ad6, and Ad7). Although BGM is the most common cell line used for ­isolation of enteric viruses from environmental samples adenoviruses will not produce CPE in this cell line although they can be detected by ICC-PCR.27,35 2.1.3.2 Antibody-Based Methods Antibody-based techniques have been developed for detecting and identifying adenoviruses in clinical samples, but have rarely been used with environmental samples. Both groupspecific techniques (e.g., detecting all human or primate adenoviruses only) and species-specific techniques (e.g., detecting Ad40 or Ad41 only) have been developed. A group-specific indirect immunofluorescence technique has been used to observe nongrowing (do not replicate or produce CPE) adenoviruses obtained from stool samples in tissue cultures.46 Only two studies have used antibody techniques for adenovirus detection in environmental samples. One used a group-specific immunofluorescence assay to detect adenoviruses in primary sludge from wastewater treatment plants.47 The viruses were visualized in HEp-2 cell cultures in which primary sludge concentrate had been added. The second study compared cell

27

culture, immunofluorescence, and in situ DNA hybridization methods for the detection of the enteric adenoviruses in raw sewage.25 While one of the cell-culture methods (using HEp-2 cells) and the immunofluorescence method (using group­specific antibodies) yielded nearly equivalent results, the average levels detected using the in situ DNA hybridization technique were approximately 40% greater. 2.1.3.3 Nucleic Acid Probes Gene probes have been developed to detect enteric adenoviruses in clinical and environmental samples, but have thus far seen limited use because they are not as sensitive or as easy to use as PCR methods. Genthe et al.48 used Ad40 and Ad41 specific digoxigenin (DIG)-labeled DNA probes for enteric adenovirus detection in both raw and treated water. Nevertheless, the viability of the adenoviruses detected using this method was questionable since they were still detectable after exposure to 20 mg/l chlorine. 2.1.3.4  PCR-Based Techniques The advent of PCR techniques has provided faster, more sensitive and more specific methods to detect adenoviruses in both clinical and environmental samples. These techniques do not demonstrate infectivity, however. Allard et al.49,50 used PCR to detect adenoviruses in untreated domestic sewage via nested PCR. Puig et al.39 compared cell culture, one-step PCR and nested PCR using sewage and river water samples. Nested PCR was found to be the most sensitive technique, allowing for the detection of < 10 particles. This is 100–1000 times more sensitive than traditional cell culture-based detection methods. Using similar techniques, Pina et al.26 were able to detect human adenoviruses in sewage, river water, seawater, and shellfish. They suggested that the detection of human adenoviruses by PCR be used as an indicator of human viral contamination of the environment. A nested multiplex PCR for detection of human enteric adenoviruses, hepatitis A virus and enteroviruses in sewage and shellfish was reported by Formiga-Cruz et al.51 The limit of detection was approximately one genome copy for adenovirus and ten copies for enterovirus and hepatitis A virus per PCR reaction using cell cultured viruses. The lower detection of enteroviruses may reflect the addition steps to perform RT-PCR for the detection of the RNA viruses. 2.1.3.5  Integrated Techniques A combination of cell culture and PCR has been used as a method to assess the viability of viruses and to increase the speed of identification (i.e., reduce the need for another passage in cell culture). In such methods, PCR is used to detect the presence of viruses growing in cell culture.51 Chapron et al.29 employed this method to detect Ad40 and Ad41 in surface water samples in BGM cells. The viruses did not produce CPE, yet could be detected by PCR. Ko et al.52 developed an RT-PCR method for the detection of Ad2 and Ad41 mRNA in cell culture. Only infectious adenoviruses are detected using this method because only viable viruses are able to produce mRNA during replication in cell culture.

28

Choo and Kim35 compared the detection of adenoviruses in oysters by ICC-PCR in BGM and human lung epithelial cells (A549) along with direct detection in the oyster samples by PCR. They found 23.6%, 50.9%, and 89.1% of all oysters positive by cell culture, ICC-PCR, and direct PCR, respectively. This suggests that not all of the adenoviruses in the oysters were viable. Rigotto et al.53 also reported the greater sensitivity of nested PCR over IC-PCR. Nested PCR was capable of detecting 1.2 plaque forming units (PFU) of Ad5 per gram of tissue vs. 120 CFU/g by ICC-PCR.

2.2  Methods Virus detection in foods usually involves an extraction step followed by a concentration step. To date the only published methods for adenovirus detection in foods have been aimed at its isolation from shellfish. Shellfish extracts and concentrates can be toxic to cell cultures used in virus detection and may have more substances than other foods which may interfere with detection by molecular methods (PCR). No method for the extract/concentration or molecular detection is 100% efficient and should not be expected to be so because of variability in the individual genomes resistance to reagents used in extraction and processing. Another issue is that the volume of the extract may be large relative to the volumes used in the molecular assay limiting the sensitivity of the assay. All of these factors should be taken into consideration when reporting results.

2.2.1 Sample Preparation Due to the low number of viruses found in most surface and ground water samples, viruses are first concentrated from volumes ranging from 10 to 1000 l. Methods commonly used are adsorption to positively or negatively charged microporous filters in a pleated cartridge format, adsorption to positively charged glass wool and ultrafiltration. Viruses are then desorbed or eluted in small volumes of liquid. Fields and Metcalf54 first reported the concentration of adenoviruses using negatively charged filters. Enriquez and Gerba19 demonstrated the use of positively and negatively charged filters for the concentration of Ad40 from tap, sea and waste waters. Jiang and Chu55 used ultrafiltration to concentrate adenoviruses from surface waters and Van Heerden et al.30 used positively charged glass wool to isolate adenoviruses from conventionally treated drinking water. Methods have only been developed for the detection of adenoviruses from shellfish. This usually involves an extraction step from the shellfish, a concentration step and then preparation for PCR. These methods are identical to what is used for other enteric viruses. Methods for used isolating enteroviruses from other foods would likely be useful for adenoviruses.56 The only consideration is that adenoviruses tend to be more sensitive to inactivation at pH levels above 9 than enteroviruses. After shucking usually several shellfish are pooled for processing. Sometimes certain organs such as the gills, digestive gland, or versa are extracted and then pooled. Since viruses may occur in the shellfish meat usually the shellfish

Molecular Detection of Foodborne Pathogens

are processed whole after removal from the shell. The shellfish meat is them homogenized in a blender or stomacher. This is followed by centrifugation to pellet the solids and virus. The supernatant is then discarded. Ultracentifugation can be used to ensure pelleting of the virus26 or the pH lowered to 5.0 and conductivity adjusted to ensure adsorption of the virus to the tissue.57 The virus is then eluted from the homogenized tissues by addition of glycine buffer at pH 9.5–10.0 or with glycine buffer at pH 7.5 and the salt concentration increased to 8000 mg/l and the solids discarded. The eluate is then concentrated further by ultracentrifugation, ultrafiltration, or acid precipitation at pH 4.5. The method reported by Pina et al.26 involving glycine buffer elution and ultracentrifugation or minor modifications appears to be the most commonly used method at present for adenoviruses.33,34,51 The following methods described the processing of shellfish sample prior the analysis by PCR for the detection of adenovirus. This method consists of the use of an alkaline glycine solution (0.25 M pH 10.00) to promote the detachment of viruses from meat. Then the sample is clarified by centrifugation and the viruses are concentrated from the elution solution by polyethylene glycol (PEG) precipitation (8% PEG and 0.3 M of NaCl). The elution of the viruses from shell fish meet was described by Formiga-Cruz et al.34 and Pina et al.26 The conditions for PEG precipitation were described by Lewis and Metcalf.58 Materials: (1) Blender (2) 500 ml Erlenmeyer flask (3) 0.25 M glycine solution: for 1 l distilled water, add 18.75 g of glycine and adjust the pH to 10.00 with 1 N NaOH (4) 250 ml conical centrifuge bottle (5) PEG 6000 (6) Sodium chloride (7) Floor swing bucked refrigerated centrifuge (8) Horizontal shaker (9) 1 N HCl solution (10) 1 N NaOH solution Virus extraction procedure: (1) Homogenize 33 g of shell fish meat with 167 ml of 0.25 M glycine solution in a blender at maximum speed for 30 sec (2) Decant the homogenize solution in a 500 ml Erlenmeyer flask and stir for 15 min (3) Adjust the pH of the solution to 7.3 with 1 N HCl solution (4) Centrifuge at 2500 × g for 30 min (5) Keep the supernatant. Measure the volume of the solution Concentration of virus by PEG precipitation: (1) For 200 ml of eluted sample, add 16 g of polyethylene glycol, and 3.5 g of NaCl (2) Decant the solution in a 250 ml conical bottle

29

Adenoviruses

(3) Put the bottle horizontally in the orbital shaker, shake at 100 revolutions/min overnight at 4°C (4) Centrifuge at 4200 rpm for 1 h (5) Add 5 ml of phosphate buffer, using the pipette, break the pellet and agitate the bottle with a voltex for 1 min (6) Measure the volume on the reconstituted concentrated sample (7) Store the sample at –20°C until further analysis

2.2.2  Detection Procedures For the nucleic acids extraction, we describe a modification of the guanidine thiocianated extraction method described by Boom et al.59 However, any commercial kit for the DNA extraction from plasma or stool sample would work. The described nested-PCR is specific for the hexon gene for all human adenovirus which includes the groups A through F. These set of primers were previously described by Avellon et al.67 Nested PCR consists of two rounds of amplification: the first amplifies the target region from the viral genome and the second round amplifies a smaller region inside the product of the first round amplification. The PCR products are analyzed by agarose gel electrophoresis. Materials: (1) Guanidine lysis buffer: 120 g guanidine thiocyanate in 100 ml TE buffer, 11 ml of 5 M NaCl, 11 ml 3 M sodium acetate (NaOAc) pH 5.5, and 3.5 ml of poplyadenylic acid 5′ potassium salt (1 mg/ml) (2) 100% proof ethanol (3) 70% ethanol solution (4) Sterile nuclease free water (5) Silica spin minicolumns (high-bind RNA minicolumns, Promega Biotek, others manufacturers columns such as Qiagen will work) (6) 2 ml collection tubes (7) 1.5 ml microcentrifuge tubes (8) 5 U/µl hot start taq polymerase (qiagen hotstart or Applied Biosystems gold Taq) (9) 10 × PCR buffer (provided with Taq polymerase) (10) 25 mM MgCl2 (provided with the Taq polymerase) (11) 25 pmoles/µl stock primers (12) 2.5 mM each dNTPs solution (10 mM total) (13) Sterile nuclease free water (14) Micropipets (0.5–10 µl, 10–100 µl, 100–1000 µl sizes) (15) Barrier sterile tips (16) 10% bleach solution (50 ml commercial bleach solution, 450 ml of water) (17) First round PCR primers are: ADHEX1F 5′-AACACCTAYGASTACATGAAC-3′ ADHEX2R 5′-KATGGGGTARAGCATGTT-3′ Fragment size of 473 bp (18) Second round PCR primers are: ADHEX2F 5′-CCCMTTYAACCACCACCG-3′ ADHEX1R 5′-ACATCCTTBCKGAAGTTCCA-3′ Fragment size of 168 bp

Nucleic acid extraction: (1) Add 100 µl of lysis buffer to a 1.5 ml microcentrifuge tubes (2) Add 100 µl of sample to the tubes and mix then with a voltex for 15 sec (3) Incubate at room temperature for 10 min (4) Remove the drops from the lip by a brief centrifugation (5) Add 200 µl of 100% proof ethanol (6) Mix with a voltex for 15 sec (7) Load the 400 µl of sample mixture into the silica minicolumns (8) Centrifuge for 1 min at 16000 × g (9) Place the column in a new collection tube (10) Add 500 µl of 70% ethanol and centrifuge at max speed for 1 min (11) Repeat the wash steps 9 and 10 one more time (12) Place the column in a collection tube. Dry the column by centrifugation at 16000 × g for 1 min (13) Put the column in a sterile 1.5-ml tube (14) Add 50 µl of sterile nuclease free water in the center of the column without touching the walls (15) Let incubate for 1 min and then centrifuge 16000 × g for 1 min (16) Keep the flow through store at –20°C for future PCR analysis Nested PCR procedure: (1) Prepare first round PCR mixture (50 µl) consisting of 1 × PCR buffer, 2.0 mM of MgCl2, 200 µM of each dNTP, 25 pmoles of each primers (ADHEX1F and ADHEX2R) for the first PCR, 1.5 U of Taq polymerase, 2 µl extracted virus DNA, and nuclease free water for a final volume of 50 µl. Prepare a master mixture for multiple reactions and adjust by increasing the volume 5% (to consider the lost of mixture during handling). Specifically, the following calculation is for a master mixture for ten reactions: (i) In a sterile 1.5 ml Eppendolf tube add: 52.5 µl of 10 ×  PCR buffer 42 µl of 25 mM MgCl2 solution 10.5 µl of Primer F (ADHEX1F) 10.5 µl of Primer R (ADHEX2R) 2.1 µl of Taq polymerase 260 µl of water (ii) Aliquot 40 µl of PCR mixture into each PCR tube and add 10 µl of sample (2) Conduct PCR amplification in a thermal cycler using the following program: one cycle of 94°C for 10 min; 35 cycles of 94°C for 30 sec, 50°C for 30 sec, and 72°C for 30 sec; one cycle of 72°C for 10 min. Use the heated lip option. (Note: If this option is not available then add a drop of mineral oil to the reaction to avoid the vaporization.) (3) Prepare second round PCR mixture (50 µl) consisting of 1 ×  PCR buffer, 2.0 mM MgCl2, 200 µM of each dNTP, 0.5 µM each primers (ADHEX2F

30

and ADHEX1R) for the second round of PCR, 1.5 U of Taq polymerase, 2 µl of first round PCR, and nuclease free water for a final volume of 50 µl. Specifically, the following calculation is for a master mixture for ten reactions: (i) In a sterile 1.5 ml eppendolf tube add: 52.5 µl of 10 ×  PCR buffer 42 µl of 25 mM MgCl2 solution 10.5 µl of nested Primer F (ADHEX2F) 10.5 µl of nested Primer R (ADHEX1R) 2.1 µl of Taq polymerase 328 µl of sterile nuclease free water (ii) Aliquot 48 µl of PCR mixture into each PCR tube and add 2 µl of first round PCR product. The cycling conditions of the second round PCR are the same used for first round PCR. (4) Agarose gel electrophoresis (i) Prepare a 2% agarose gel in 0.5 × TBE in a 250-ml Erlenmeyer flask adding 2 g agarose and 100 ml 0.5 × TBE buffer (ii) Heat the gel in a microwave oven until the ­agarose dissolves. Add 5 µl of 10 mg/ml ethidium bromide stock solution (50 µg/ml working solution) after the agarose cools down (~45°C) and before pouring. (iii) Pour the gel slowly, trying to reduce the formation of bubbles. Put the well comb in its correct position. Let the gel stand for at least 30 min. (iv) Locate the gel wells in the negative side of the electrophoresis chamber. Add 0.5 × TBE buffer until the lever reaches more than 2 mm above the gel. (v) Mix 10 µl of the sample with 2 µl of 6 × loading buffer. (vi) Using a pipette with a fine tip, load the sample into the well. Load at step DNA ladder every six samples (amount recommended by manufacturer). (vii) Check that the polarity is in the correct orientation (well in the negative side, running to positive side). (viii) Applied as follows: 5 V per cm gel length. (ix) Stop the electrophoresis when it reached 70% of the gel length. Use as reference the faster dye of the loading buffer. (x) Visualize the PCR products using a UV transilluminator. (xi) The PCR product size is estimated by comparing it with the step ladder and with the positive control. The 100 bp step ladder has step increases from 100 bp to 1000 bp. A positive sample for adenovirus should show a band between 100 bp and 200 bp with 168 bp. Also, a positive sample should have the same running distance as the positive control.

Molecular Detection of Foodborne Pathogens

Comment on quality control: (1) All the areas for the analysis should be separate in different rooms: one room exclusive for mixing PCR reagents, one room for handling the samples, one for the nested PCR, and another for gel electrophoresis. (2) We recommend using a different PCR workstation/ hood with UV lamp for preparing the master mix and another for the addition of the first round PCR product to the second round PCR mixture. Use a biological hood type 2 for handling the samples. Before and after uses, the hoods should be cleaned with 10% bleach solution and turn on the UV light for 30 min. The bleach can be inactivated with 2% sodium thiosulfate solution and washed with water. (3) Open the reagents only inside the workstation, and the samples and PCR products are only opened in their respective workstation. (4) Keep the equipment in each respective room and not used then in other areas (i.e., pipets, tips, and different lab coats are exclusively used in each room). (5) The PCR product is only opened in the workstation for samples and in the electrophoresis room (negative pressure from the main laboratory).

2.3 Conclusions and Future Perspectives Adenoviruses, like the enteroviruses, cause a wide range of illness many of which are not thought of as being food or waterborne. This, combined with the long time needed to produce cytopathogenic effects in cell culture previously resulted in few studies on their occurrence in the environment and role in food and waterborne diseases. While waterborne transmission is well documented, the role of food in their transmission is unknown. Certainly documenting their presence in food, especially shellfish suggests foodborne transmission is possible. Human adenoviruses are more common in sewage contaminated waters and it has been suggested by several groups of investigators that they may be useful indicators of other enteric viruses in water and shellfish. Adenoviruses also have the added advantage of unlike most human enteric viruses, they have a DNA genome, eliminating the need for RT-PCR. However, recent research on the occurrence of adenoviruses in areas remote from human sewage contamination suggests that other sources besides humans or exceptionally long survival times in the environment.61 Clearly additional research is needed to assess the usefulness of human adenoviruses as indicators of human viral contamination. The application of PCR makes possible for the first time low cost and simple methods for the detection of adenoviruses in food and water. This will help us better understand the role of these vehicles in their transmission. Currently all molecular methods suffer from inability to determine viability without the use of cell culture, small assay volumes, quantification at low numbers of genome copies, interference with substances in concentrates, and the loss of virus during sample processing. These are major challenges that need to

Adenoviruses

be overcome to take full advance of molecular approaches for virus detection in foods.

References





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31 22. Villena, C. et al. A large infantile gastroenteritis outbreak in Albania caused by multiple emerging rotavirus genotypes. Epidemiol. Infect., 131, 1105, 2003. 23. Divizia, M. et al. Waterborne gastroenteritis outbreak in Albania. Water Sci. Technol., 50, 57, 2004. 24. Irving, L.G. and Smith, F.A. One-year survey of enteroviruses, adenoviruses, and reoviruses isolated from effluent at an activated-sludge purification plant. Appl. Environ. Microbiol., 41, 51, 1981. 25. Hurst, C.J., McClellan, K.A. and Benton, W.H. Comparison of cytopathogenicity, immunofluorescence and in situ DNA hybridization as methods for the detection of adenoviruses. Water Res., 22, 1547, 1988. 26. Pina S. et al. Viral pollution in the environment and in shellfish: human adenovirus detection by PCR as an index of human viruses. Appl. Environ. Microbiol., 64, 3376, 1998. 27. Rodriguez, R.A., Gundy, P.M. and Gerba, C.P. Comparison of BGM and PLC/PRC/5 cell lines for total culturable viral assay of treated sewage. Appl. Environ. Microbiol., 74, 2583, 2008. 28. Krikelis, V. et al. Seasonal distribution of enteroviruses and adenoviruses in domestic sewage. Can. J. Microbiol., 31, 24, 1985. 29. Chapron, C.D. et al. Detection of astroviruses, enteroviruses, and adenovirus types 40 and 41 in surface water collected and evaluated by the information collection rule and an integrated cell culture-nested PCR procedure. Appl. Environ. Microbiol., 66, 2529, 2000. 30. Van Heerden, J. et al. Incidence of adenoviruses in raw and treated water. Water Res., 37, 3704, 2003. 31. Lee, H.K. and Jeong, Y.S. Comparison of total culturable virus assay and multiplex integrated cell culture-PCR for reliability of waterborne virus detection. Appl. Environ. Microbiol., 70, 3632, 2004. 32. Enriquez, C.E. and Gerba, C.P. Concentration of enteric adenovirus 40 from tap, sea, and waster water. Water Res., 95, 2554, 1995. 33. Munianin-Mujika, I. et al. Comparative analysis of viral pathogens and potential indicators in shellfish. Int. J. Food Microbiol., 83, 75, 2003. 34. Formiga-Cruz, M. et al. Distribution of human virus contamination in shellfish from different growing areas in Greece, Spain, Sweden, and the United Kingdom. Appl. Environ. Microbiol., 68, 5990, 2002. 35. Choo, Y.J. and Kim, S.J. Detection of human adenoviruses and Enteroviruses in Korean oysters using cell culture, integrated cell culture-PCR, and direct PCR. J. Microbiol., 44, 1662, 2006. 36. Umesha, K.R. et al. Prevalence of human pathogenic enteric viruses in bivalue molluscan shellfish and cultured shrimp in south west coast of India. Int. J. Food Microbiol., 122, 279, 2008. 37. Enriquez, C. E., Hurst, C.J. and Gerba, C.P. Survival of the enteric adenoviruses 40 and 41 in tap, sea, and waste water. Water Res., 29, 2548, 1995. 38. Hernroth, B. and Allard, A. The persistence of infectious adenovirus (type 35) in mussels (Mytilus edulis) and oysters (Ostrea edulis). Int. J. Food Microbiol., 113, 296, 2007. 39. Puig, M. et al. Detection of adenoviruses and enteroviruses in polluted waters by nested PCR amplification. Appl. Environ. Microbiol., 60, 2963, 1994. 40. Echavarria, M. et al. PCR method for detection of adenovirus in urine of healthy and human immunodeficiency virusinfected individuals. J. Clin. Microbiol., 36, 3323, 1998.

32 41. Tani, N. et al. Seasonal distribution of adenoviruses, ­enteroviruses and reoviruses in urban river water. Microbiol. Immunol., 39, 577, 1995. 42. Brown, M. Laboratory identification of adenoviruses associated with gastroenteritis in Canada from 1983 to 1986. J. Clin. Microbiol., 28, 1525, 1990. 43. Grabow, W.O., Puttergill, D.L. and Bosch, A. Propagation of adenovirus types 40 and 41 in the PCL/PRF/5 primary liver carcinoma cell line. J. Virol. Methods, 37, 201, 1992. 44. Pintó, R.M. et al. Detection of fastidious infectious enteric viruses in water. Environ. Sci. Technol., 29, 2636, 1995. 45. Takiff, H.E., Straus, S.E. and Garon, C.F. Propagation and in vitro studies of previously non-culturable enteral adenoviruses in 293 cells. Lancet, 2, 832, 1981. 46. Retter, M. et al. Enteric adenoviruses: detection, replication and significance. J. Clin. Microbiol., 10, 574, 1979. 47. Williams, F.P. and Hurst, C.J. Detection of environmental viruses in sludge: enhancement of enterovirus plaque assay titers with 5-iodo-2′-deoxyuridine and comparison to adenovirus and coliphage titers. Water Res., 22, 847, 1988. 48. Genthe, B. et al. Detection of enteric adenoviruses in South African water using gene probes. Water Sci. Technol., 31, 345, 1995. 49. Allard, A. et al. Polymerase chain reaction for detection of adenovirus in stool samples. .J Clin. Microbiol., 28, 2659, 1990. 50. Formiga-Cruz, M. et al. Nested multiplex PCR assay for detection of human enteric viruses in shellfish and sewage. J. Virol. Methods, 125, 111, 2005. 51. Reynolds, K.A., Gerba, C.P. and Pepper, I.L. Detection of infectious enteroviruses by an intergrated cell culture-PCR procedure. Appl. Environ. Microbiol., 62, 1424, 1996.

Molecular Detection of Foodborne Pathogens 52. Ko, G., Cromeans, T.L. and Sobsey, M.D. UV inactivation of adenovirus type 41 measured by cell culture mRNA RT-PCR. Water Res., 39, 3643, 2005. 53. Rigotto, C. et al. Detection of adenoviruses in shellfish by means of conventional-PCR, nested-PCR, and integrated cell culture PCR (ICC/PCR). Water Res., 39, 297, 2005. 54. Fields, H.A., and Metcalf, T.G. Concentration of adenovirus from seawater. Water Res., 9, 357, 1975. 55. Jiang, S., and Chu, W. PCR detention of pathogenic viruses in southern California urban rivers. J. Appl. Microbiol., 97, 17, 2004. 56. Goyal, S.M. Methods of virus detection in foods. In Viruses in Foods, Goyal, S.M. (Ed.). pp. 101–119. Springer, New York, NY, 2006. 57. Sobsey, M.D., Carrick, R.J. and Jensen, H.R. Improved methods enteric viruses in oysters. Appl. Environ. Mirobiol., 36, 121, 1978. 58. Lewis G.D. and Metcalf, T.G. Polyethylene glycol precipitation for recovery of pathogenic viruses, including hepatitis A virus and human rotavirus, from oyster, water, and sediment samples. Appl. Environ. Microbiol., 54:1983, 1988. 59. Boom, R., C.J.A. et al. Rapid and simple method for purification of nucleic acids. J. Clin. Microbiol., 28, 495, 1990. 60. Avellón, A.P. et al. Rapid and sensitive diagnosis of human adenovirus infections by a generic polymerase chain reaction. J. Virol. Methods, 92, 113, 2001. 61. Lipp, E.R., Futch, J.C. and Griffin, D.W. Analysis of multiple enteric viral targets as sewage markers in coral reefs. Mar. Pollut. Bull., 54, 1897, 2007.

3 Astroviruses

Edina Meleg and Ferenc Jakab University of Pécs

Contents 3.1 Introduction......................................................................................................................................................................... 33 3.1.1 History, Virion Structure and Classification............................................................................................................ 33 3.1.1.1 History....................................................................................................................................................... 33 3.1.1.2 Genome Structure..................................................................................................................................... 34 3.1.1.3 Classification............................................................................................................................................. 35 3.1.2 Pathogenesis and Pathology..................................................................................................................................... 35 3.1.2.1 Physical Features....................................................................................................................................... 35 3.1.2.2 Propagation of Human Astroviruses......................................................................................................... 35 3.1.2.3 Propagation of Animal Astroviruses........................................................................................................ 36 3.1.2.4 Transmission............................................................................................................................................. 36 3.1.2.5 Pathogenesis.............................................................................................................................................. 36 3.1.3 Clinical Features . ................................................................................................................................................... 36 3.1.3.1 Characteristics of Human Illness.............................................................................................................. 36 3.1.3.2 Role in Immunocompromised Hosts......................................................................................................... 37 3.1.3.3 Immunity................................................................................................................................................... 37 3.1.4 Epidemiology........................................................................................................................................................... 38 3.1.4.1 Role in Disease.......................................................................................................................................... 38 3.1.4.2 Antibody Acquisition and Prevalence....................................................................................................... 38 3.1.5 Laboratory Diagnosis............................................................................................................................................... 38 3.1.6 Treatment and Prevention........................................................................................................................................ 40 3.2 Methods............................................................................................................................................................................... 40 3.2.1 Sample Preparation ................................................................................................................................................. 40 3.2.3 Detection Procedures............................................................................................................................................... 40 3.3 Conclusions and Future Perspectives.................................................................................................................................. 43 References.................................................................................................................................................................................... 44

3.1  Introduction

3.1.1 History, Virion Structure and Classification

Astroviruses are the members of Astroviridae family, which include both human and animal nonenveloped viruses possessing a plus-sense, ssRNA genome. In humans, astroviruses mainly produce gastroenteritis together with a broad spectrum of symptoms such as malaise, vomiting, diarrhea, fever and abdominal pain. Besides being one of the most common causes of viral gastroenteritis in young children in developed countries, astrovirus is the culprit for viral diarrhea in young children in some other parts of the world.1 Although foodborne illness resulting from viral infections is a large and growing public health problem, most countries do not have good reporting systems; therefore realistic estimation of the true burden of foodborne diseases is difficult. Based on a recent study, astroviruses account for less than 1% of foodborne illnesses in the United States.2

3.1.1.1 History Astroviruses were first identified by Madeley and Cosgrove in 1975 in the feces of hospitalized infants with diarrhea.3 Based on direct electron microscopy (EM) studies of fecal samples, astroviruses were observed as 28–30 nm particles in diameter with a distinctive five-six pointed star-like surface (Figure 3.1). This morphology distinguished astroviruses from other small, round viruses with similar size, such as picornaviruses and caliciviruses. In some preparations, bridging structures, which may be surface extensions of the virus, have been observed between adjacent astrovirus particles.4 The term “astrovirus” was named after the star morphology (astron = star; Greek), although this surface structure can only be identified in approximately 10% of the particles by EM. Interestingly, particles isolated from infected rhesus

33

34

Molecular Detection of Foodborne Pathogens

5´UTR Non-structural proteins

3´UTR

?

A(n) Structural proteins ORF 1a

ORF 1b

ORF 2

Frameshift site

Figure 3.2  Genome arrangement of astroviruses. www.tulane. edu/~dmsander/WWW/335/Diarrhoea1.gif 50 nm

Figure 3.1  Image of human astroviruses by electron microscopy. http://www.virology.net/Big_Virology/BVRNAastro.html

monkey kidney epithelial (LLCMK2) cells lack the star-like morphology, but it can be induced by a brief exposure to pH = 10 environment. Later, based on high-resolution EM5 and electron cryomicroscopy studies astroviruses seemed to be icosahedral particles with spikes or knob-like projections, and 41–43 nm in total diameter. Studies on purified cell culture-adopted HAstV-1 particles evaluated by cryo-electron microscopy and image analysis revealed a rippled solid capsid shell structure (330 Å in diameter) decorated with 30 dimeric spikes extending 50 Å from the virion surface.6 These particles, however, have not been characterized by protein composition, which is a key factor for the virus-specific infectivity and may have an effect on virion structure.7,8 A few months before naming astroviruses, in 1975 Appleton and Higgins9 reported an outbreak of mild diarrhea and vomiting among infants in a maternity ward. In their study, astroviruses particles were 29–30 nm in diameter, and did not display the special surface features, however by EM these viruses were distinct in size and morphology from the previously identified Norwalk viruses and rotaviruses. One year later specific immunologic reagents proved that these viruses were really astroviruses.10 Subsequently, viral particles, that were similar size and had the star-like surface features, were observed in gastroenteritis cases in several young mammals and birds, including mice,11 kittens,12 dogs,13,14 lambs,15 calves,16 deer,17 piglets,18 minks19 as well as turkeys.20 Gough et al.21 observed fatal hepatitis in ducklings due to astrovirus infection, and virus particles were found in liver of these animals in addition to feces. Astrovirus appears to cause species-specific infections.22 In 1981, Lee and Kurtz23 published the successful isolation of human astroviruses in the presence of trypsin in human embryonic kidney (HEK) cells and passage in LLCMK2 cells, which definitely distinguished them from noncultivatable small, round viruses, such as caliciviruses. In the late 1980s, Herrmann et al.24,25 developed an enzyme immunoassay (EIA) that detects viral antigen. Although, the virus was well known since 1975, the real medical importance was only recognized in 1991.26 Reverse-transcription polymerase chain reaction (RT-PCR) assays were first use in 1995 to detect human astroviruses. These techniques contributed

to more detailed characterization of astrovirus strains by the analysis of nucleotide sequence information.27–29 In the past few years spreading of RT-PCR technique could be observed in studies of the prevalence of astrovirus among children with diarrhea to detect and genotype strains. 3.1.1.2 Genome Structure Astroviruses have a plus-sense, single-stranded RNA genome, which is approximately 6800 nucleotides (nt) (varies from 6.4 to 7.3 kb) in length. It is polyadenylated at the 3′ end, and surrounded by an isocahedral capsid. The genomial RNA includes 5′ and 3′ nontranslated region (UTR), and three open reading frames (ORFs), each encoding polyprotein that is proteolytically processed to yield smaller proteins (Figure  3.2). The two ORF located toward the 5′ end of the genome, designated ORF 1a (~2700 nt) and ORF 1b (~1550 nt), encode nonstructural proteins, such as an RNAdependent RNA polymerase and a 3C-like serine protease that are involved in RNA transcription and replication. ORF 1a also encodes overlapping immunogenic epitopes that are recognized by antibodies produced to intact astroviruses.30 An overlap of 60–70 nt is found between ORF 1a and ORF 1b of mammalian astroviruses (this region is only 12–45 nt long in avian viruses). This overlapping region contains signals which are essential for translation of the viral RNA polymerase through a frameshift mechanism.31,32 This region is completely conserved among human and animal astroviruses in two characteristics: a heptameric AAAAAAC sequence, and the potential to form a downstream stem-loop structure. These features are critical for the ribosomal-1 frameshifting event during the translation of the genome.31,32 The third ORF is located at the 3′ one-third of the genome, designated ORF 2, which has the greatest sequence variability in the astrovirus genome, and encodes the 90 kDa protein that is the precursor of the three capsid proteins that have been described for human astroviruses.33,34 Structural proteins encoded by ORF 2 are translated from the so-called subgenomic (sg) RNA.33,34 The more conserved amino-terminal region of the astrovirus capsid protein has an important function in assembly of the capsid core, while the hypervariable carboxy-terminal form the spikes of the virion and participate in the early interactions between the virus and the host cells.35 ORF 1b and ORF 2 overlap in eight nt. The 3′ UTR of HAstV genome, which is located between ORF 2 and the polyA tail is 80–85 nt long (this sequence can be longer [130–305 nt] in avian viruses).36

35

Astroviruses

The final 19 nt of ORF 2 and the 3′ UTR are thought to be important for interacting with the viral RNA replicase and cellular proteins. These regions are highly conserved among all known HAstV serotypes.37 During infection of susceptible cells, two RNA species have been observed: the full-length genomic RNA (gRNA), and an sgRNA (~2.4 kb in length).38,39 Both RNA species are initially observed at 12 hours postinfection in LLCMK2 cells.39 Synthesis of the negative-sense RNA of astrovirus has not been well studied. By all means, the gRNA is probably a template to synthesize the full-length negative-sense RNA, which is a template to produce both the full-length gRNA and the sgRNA. The synthesis of sgRNA probably requires an internal sequence in the full-length negative sense RNA to serve as promoter for the virus transcriptase. However, the identity of this promoter in astrovirus has not been defined; it is thought that about 120 nt of the ORF 2 region might be an important sequence of the promoter.40 Part of this region includes the sequence AUUUGGAGNGGNACCNAAN5-8 AUGNC (the ORF 2 start codon is underlined; N can be any of the four nucleotides), which is highly conserved among all members of Astroviridae.41 After astrovirus entry, the gRNA is used as a template to synthesize the virus nonstructural proteins. The primary protein product coded by ORF 2 is observed abundantly 12 hours after infection39 (nonstructural proteins are initially detected 6 hours postinfection42). 3.1.1.3  Classification Genome arrangement of astroviruses is similar to the genome of Picornaviridae and Caliciviridae,43 but the size, number, and processing of polyproteins, the lack of an RNA-helicase domain in astroviruses, and the use of a ribosomal frameshifting mechanism distinguish them from similar viruses. Therefore astroviruses are classified into a separate family, the Astroviridae.44 Astroviruses have been isolated from both humans and several animal species. According to the origin of the virus and the genome structure, two genera have been distinguished within the family: mamastrovirus (infect mammals) and avastrovirus (including viruses from avian species). Viruses in the genus mamastrovirus are more closely related to each other than those viruses within the avastrovirus genus.36,45 Serologic relatedness between viruses isolated from different species, even within the same genus has not been identified.46,47 Human astroviruses have been grouped into eight serotypes (HAstV-1 to HAstV-8) according to immunfluorescence and neutralization assays, as well as immunelectron microscopy that use hyperimmune sera to raise different culture adapted strains.23,48–52 Common epitopes of the capsid protein in cases of all known HAstV serotypes were identified. These epitopes are widely used in many different diagnostic assays.24,25 Recently, more sensitive molecular methods are used, that enabled the classification of human astroviruses on the basis of the sequence similarity of specific genome regions. Nowadays, different regions of the genome, obtained

by RT-PCR, have been widely used to group HAstVs into different genotypes; though depending on the specific region of the analyzed genome (ORF 1a, 1b or ORF 2), strains can be grouped differently.53 Several animal serotypes of astroviruses have been identified. At least two, but probably three serotypes are found among bovine astroviruses (BAstVs) based on the crossreactivity to specific sera. Two serotypes have been identified among turkey (TAstVs)36 and chicken astroviruses (CAstVs).46 Many serotypes of astroviruses have been identified in consequence of different serological studies, suggesting that additional types of astrovirus might exist.47,54

3.1.2  Pathogenesis and Pathology 3.1.2.1  Physical Features Astroviruses are one of the most resistant viruses. They show resistance against different physical and chemical agents, such as chloroform, a variety of nonionic, anionic as well as zwitterionic detergents, even lipid solvents; they are able to maintain their infectivity at 60°C for 5 min (HAstV)5,22 or 10 min (TAstV), and at ultralow temperature (–70 to –85°) for 6–10 years, but repeated freezing and thawing is detrimental55 to them, particles are resistant to treatment from pH 3 to 10.5,22 The extreme stability of astroviruses against environmental factors suggests that traditional pasteurization procedures cannot completely inactivate them. Furthermore, astroviruses are able to persist under severe environmental conditions, they endure on inanimate surfaces, on human hands, in dried human and animal fecal materials, in water, on kitchen surfaces, food preparation areas, hospital as well as cruise ship cafeterias, on carpets and hospital lockers.56 There is a lack of information on the survival of astrovirus on foods. Information is also lacking on the efficiency of current washing and decontamination procedures for the removal of astrovirus. It was already demonstrated that traditional disinfection procedures do not eliminate astroviruses by water treatment. 3.1.2.2  Propagation of Human Astroviruses Serial passage of astrovirus in HEK cells requires incorporation of 10 μg/ml trypsin in the serum-free growth media.23 Higher levels of typsin do not improve the viral yield, while lower levels (for example the 0.5 μg/ml necessary for rotavirus growth) are not sufficient to maintain astrovirus in cell culture. After several passages of HEK-293 cells, viruses were able to grow in primary baboon kidney (PBK) cells and in a continuous cell line of LLCMK2 cells. Attempts to establish serial passage by direct inoculation of PBK or LLCMK2 cells with fecally derived astrovirus were unsuccessful. Willcocks et al.57 could propagate the virus in a continuous cell line of human colon adenocarcinoma (CaCo-2) cells using 5 μg/ml trypsin in the culture medium. With this method, the first cytopathic effect (CPE) appeared after 2 days of infection. Nowadays, this cell line is commonly used to grow wild-type astrovirus strains.

36

However, adenocarcinoma cell lines (CaCo-2, T-84, HT-29, SK-CO-1 cells) are the most efficient cells to isolate HAstV, the virus are also able to grow in human liver hepatoma cells (PLC/PRF/5) and in monkey kidney-derived (MA104, Cos-1, vero) cells as well.52,58–60 CaCo-2, T-84 and PLC/PRF/5 cell lines are the most efficient to directly isolate HAstV strains from fecal suspensions.52,58–60 Adaptation of field HAstV strains to grow in cell culture has low efficiency because different astrovirus strains have different susceptibility to trypsin. Although, 10 μg/ml of trypsin has been regularly used to activate virus infectivity; recent isolation of HAstV type-8 in CaCo-2 cells required much higher (200–400 μg/ml) trypsin concentration.8 In consequence, the optimal concentration of trypsin to obtain completely activated virus can depend on the HAstV strain. 3.1.2.3  Propagation of Animal Astroviruses To date bovine,61 porcine62 and avian astroviruses have been propagated in cell culture. Bovine astrovirus was isolated by Aroonprasert et al. in primary neonatal bovine kidney (NBK) cells in the presence of 50 μg/ml trypsin. This amount of trypsin in the culture media was necessary both for direct isolation from fecal samples and for serial passage of the virus in NBK cells. Primary bovine embryo kidney (BEK) cells were also efficient in growing bovine astroviruses using feces and cell culture adopted viruses as the starting material.61 A porcine astrovirus was isolated in embryonic swine kidney cells using 50 μg/ml trypsin in the culture medium. Fecal samples from infected pigs were filtrated and used directly for infection. However, maximal CPE was evident 4–5 days after inoculation of cell monolayers, the CPE was not formed if trypsin was removed from the medium or if the virus inoculum was mixed with convalescent phase serum.62 Chicken astroviruses were successfully adapted to both chicken embryo liver (CEL) and chicken hepatocellular carcinoma (LMH) cells. Tracheal swabs, lymphocytes, and intestine homogenates from infected chickens were particularly used as a source of the virus. Duck astroviruses were also adapted to these cells (CEL and LMH) without the addition of trypsin to the medium.46 Adaptation of turkey astrovirus to avian (as turkey and chicken embryo-derived cells) and mammalian (e.g. CaCo-2, vero) cell lines was unsuccessful, although they could be replicated in turkey embrionated eggs.63–65 Astroviruses from other vertebrates, like lambs,15 red deer,17 cats26 and dogs13,14 have also been isolated in cell cultures, but serial passage of the virus has not been performed. 3.1.2.4 Transmission Astrovirus can be transmitted through the fecal-oral route, by person-to-person contact, from fecally contaminated fingers to foods or to work surfaces and door handles. There is a significant risk of contamination from field workers who do not have adequate on-site toilet and hand-washing facilities.56 Astroviruses can be disseminated over a wide area in aerosol droplets (produced by vomiting), which is a particular hazard for exposed food or surfaces with subsequent transfer to foods.

Molecular Detection of Foodborne Pathogens

Astroviruses can be transferred with contaminated food and water from different origins.52,66–70 Sequence analysis of HAstV strains detected from both clinical samples and water supplies verified that water could be an important source for HAstV contamination, because virus strains from both origins were identical, at least in the specific genome region analyzed.69 It was shown, that poliovirus can infiltrate into the roots and body of plants from the soil,71 and consequently it is probable, that astroviruses also have the same feature. 3.1.2.5  Pathogenesis Pathogenesis of human astrovirus infections has been extensively studied. Recent histopathologic examinations show that astrovirus infects the mature epithelial cells of the small intestine, especially in the jejunum and in the duodenum.72 Severe diarrhea caused by villus atrophy in the intestine suggests that the inflammatory response does not play an important role in the pathogenesis of astrovirus.72 Other mammalian astroviruses can infect epithelial cells (OAstV, BAstV), subepithelial macrophages (OAstV), as well as M cells (BAstV) of the small intestine.73,74 OAstV particles were also observed in vacuoles of the enterocytes.73 OAstV infection was characterized by transient villus atrophy and crypt hypertrophy, which resulted in severe diarrhea after 2–4 days of infection. BAstV was unable to induce diarrhea in gnotobiotic animals, nevertheless, inflammatory mononuclear cells above the dome villi were observed on infection with this virus.74 In addition, the lamina propria was infiltrated with neutrophils and cells with degenerate nuclei were present. Lymphoid cell depletion was noted in the central region of germinal centers beneath the infected dome villi. In the case of turkey astrovirus 2 (TAstV-2) infection, mild crypt hyperplasia was observed after 1 day of infection in the proximal jejunum, while after 3–5 days of infection the same manifestation was observed in the distal jejunum and ileum, as well as in the duodenum.75 Electron microscopy studies revealed intracytoplasmic astrovirus aggregates in enterocytes on the sides and base of villi in the ileum and distal jejunum on day 3 postinfection. Astrovirus infection do not cause inflammation in humans72 and turkeys,65 but induce apoptosis in cultured cells,76 which suggest that this form of programmed cell death, could contribute to diarrhea in some species. Several other mechanisms could also contribute to the gastroenteritis due to astroviruses.

3.1.3 Clinical Features 3.1.3.1  Characteristics of Human Illness HAstV are the causative agents of viral gastroenteritis worldwide mainly in children (under the age of 5). Within the four childhood gastroenteritis virus (rotavirus, enteric adenovirus, astrovirus and calicivirus) HAstV is the second most common viral agent that causes diarrhea in young children evaluated in outpatient settings.26 Astrovirus infections have also been recognized in elderly, institutionalized patients77,78 and

37

Astroviruses

immunocompromised individuals. HAstV type 1 has been detected as the predominant strain in most countries.22,51 However, the incubation period in most HAstV infections is 3–4 days; a shorter incubation time of 24–36 hours was also documented during an outbreak of gastroenteritis in a Japanese kindergarten.79–81 Generally, human astroviruses induce a mild, watery diarrhea that typically lasts for 2–3 days, associated primarily with vomiting, fever, anorexia, abdominal pain and a variety of constitutional symptoms lasting no more than 4 days.22,82 Dehydration also can occur in patients with underlying gastrointestinal disease, poor nutritional status, or mixed infections83 (Table 3.1). Prolonged lactose intolerance and sensitivity to cow’s milk have been described.84,85 Persistent gastroenteritis due to astrovirus has been associated with serotype 3.86 Deaths related to astrovirus infection are extremely rare, although have been reported.87 Severe intussusception caused by HAstV infection was also documented in a child hospitalized with gastroenteritiss.88 In an Argentinean outpatient study with children under 36 months of age, astrovirus was associated with 12.4% of the diarrhea episodes; fever was present in 41.6%, and 16.7% of the patients required hospitalization.89 In Egypt among children under the age of 3 years, the total incidence of diarrhea due to astrovirus was equal to rotavirus; and severe dehydration arose out of 17% of astrovirus infected patients.90 Astroviruses cause infection at relatively low doses in humans, and appear in food which are usually obtainable in easy-to-use form, and therefore are not subjected to cooking conditions that kill them. Infected food workers may shed virus for longer periods of time, and for that reason may remain infectious even after full recovery.92 In children it may be difficult to distinguish diarrhea caused by astrovirus from that caused by rotavirus on clinical grounds alone.22,26 However, in general astrovirus diarrhea is less severe when compared to symptomatic rotavirus infection, as it does not cause significant dehydration and patients are less likely to require hospitalization.26,45,85 3.1.3.2 Role in Immunocompromised Hosts HAstVs cause chronic diarrhea among immunosuppressed patients in all age groups. HAstVs cause infection more frequently in patients with several immune diseases, such as chronic lymphocytic leukemia, congenital T-cell immunodeficiency, human immunodeficiency, combined immunodeficiency, Waldenstrom’s macroglobulinaemia and immunodeficiency polyendocrinopathy.93–96 Depletion of CD4 + T-cells by disease or iatrogenic means (for example chemotherapy) develops prolonged astrovirus diarrhea.93,97,98 Among HIV-infected patients, several viruses (for example astroviruses, adenoviruses, picobirnaviruses) were found more often in the stools of those with diarrhea (n = 65) than those without diarrhea (n = 65).95 HAstVs have been associated with outbreaks in bone marrow transplant patients.99 Chronic astrovirus diarrhea has been published in a child, who received a bone marrow transplant for combined immunodeficiency.22 The infection persisted until the child’s death, but no antibodies to astroviruses were detected in the serum.

Table 3.1 Clinical Symptoms Associated with Human Astrovirus Infection Diarrhea

Incidence 72–100% Duration 2–3 days (average) Maximum number of stools 4/24 hours Incidence of bloody diarrhea 0% Abdominal pain Incidence 50% Vomiting Incidence 20–70% Duration 1 day (average) Maximum number of vomiting 1/24 hours Fever Incidence 20–25% Maximum 37.9°C Dehydration Incidence to a degree 24–30% Incidence of severe dehydration 0–5% Hospitalization Incidence 6% Duration 6 days (average) Bronchiolitis Incidence 33% Otitis Incidence 13% Severity score (1–20)* 5 (average) Admission diagnosis of gastroenteritis 18.7–48% Source: Adopted from Walter, J.E., Mitchell, D.K., Curr. Opin. Infect. Dis., 16, 247, 2003. *20 points scoring system according to Ruuska and Vesikari.91

3.1.3.3  Immunity At present the determining factors of immunity to astrovirus are not well understood. Astroviruses primarily infect two age groups (young children and the elderly) and institutionalized patients. The age distribution of symptomatic infection suggests that antibodies to astrovirus acquired in childhood provide a certain protection from illness through adult life and that immunity decreases late in life. Studies revealed that in volunteers with detectable serum astrovirus antibody, diarrhea did not manifest clinically after virus challenge.79 Indirect evidence suggests that astrovirus-specific antibodies play a role in limiting infection in the host. Gamma globulin pools in the USA and Japan contain antibodies to human astroviruses, suggesting that astrovirus infection is common.81,100 Studies in the UK have shown that antibodies to astrovirus are acquired in early childhood; 70% have antibody by school age, 75% by 10 years of age, and 77% by early adulthood.101 The normal mucosal immune system is important in the protection of individuals from repeated human astrovirus infections.102 CD4 + T-cells that recognize human astrovirus antigens in a human leukocyte antigen (HLA)-restricted manner have been found in the lamina propria of intestinal tissue of healthy adults.102 Upon activation, these human astrovirus CD4 + T-cells may play a role in preventing repeated astrovirus infections by production of helper T-cell subtype 1-type cytokines, interferon gamma and tumor necrosis factor, providing a defense barrier at the portal of entry.

38

The role of the humoral immune response in animals to restrict astrovirus infection is not clear. It was demonstrated, that virus replication in small turkeys infected with TAstV was limited, however the infection did not induce significant adaptive immune response. No protection was observed against TAstV on secondary challenge; and the restricted virus replication was attributed to an inherent response, cured by production of nitric oxide.103

3.1.4 Epidemiology Human astrovirus infections have been detected worldwide, principally in young children suffering from diarrhea.9,77,95,104,105 Human astroviruses cause disease in (i) infants and young children, (ii) elderly institutionalized patients, (iii) immunocompromised hosts, and (iv) otherwise healthy individuals that come into contact with ­astrovirus-contaminated food or water. Studies in Australia,106 Thailand26 and Guatemala83 have revealed HAstV as the second most common cause of gastroenteritis in children, after rotavirus, with incidences varying from 4.2% to 8.6%. Large outbreaks caused by astroviruses through contaminated food, which affect thousands of persons in Japan, have been reported among otherwise normal school-age children and adults.107,108 Age distribution of HAstV can vary. In a Spanish study, 80% of children under 3 years of age were infected with astroviruses.109 In Egypt age-specific HAstV and rotavirus incidences were similar (0.38 for infants under 6 months, 0.40 for infants between 6 and 11 months, and 0.16 for children 12–23 months).90 Astrovirus infections occur primarily in the winter months in temperate regions and in the rainy seasons in more tropical climates.45,83 HAstV type 1 has been detected as the predominant strain in most countries,48,69,106,110 and the circulation of other types in a given period is probably less frequent throughout the year in any single geographic area, but the most common serotype can vary with time and location. Hence, in the UK 72% of the community-acquired astroviruses, detected between 1975 and 1987 were serotype 1,22 while in Australia serotypes 1, 3, and 4 were most frequently found in an 18-year period.111 In Mexico, astrovirus type 2 was the predominant strain (35%).112 Although in a study in 2004, where samples were analyzed from different regions of Mexico, types 1, 2, 3, 4, 6, 7, and 8 were found, and in a region, frequency of serotype 8 and type 1 was the same.113 Both community-acquired26,85 and nosocomial9,84 infections have been described. 3.1.4.1 Role in Disease Until recently it was complicated to determine the real incidence of astrovirus infection and its role in disease, partly due to the frequency of coinfections with other pathogens.45 Development of more sensitive detection methods has clarified their role in gastroenteritis in different populations. Astroviruses frequently cause outbreaks of diarrhea in child care centers, and children less than 36 months of age attending child care settings are at the greatest risk of developing

Molecular Detection of Foodborne Pathogens

diarrhea.114 In several parts of the world astroviruses are known to be the main cause of viral diarrhea in young children.1 Astroviruses were identified as a foodborne pathogen only within the last 30 years, though they have probably been causing foodborne infections for centuries. Astroviruses are estimated to participate in less than 1% of all foodborne illness, similar to rotaviruses,2 which mean 39000 cases per year out of 3.9 million total cases due to viruses.2 Although, weaknesses and variations in foodborne disease surveillance systems (where such programmes exist at all), make the global estimation of foodborne diseases difficult. Symptoms of astroviral foodborne infection range from mild gastroenteritis to severe life-threatening syndromes. The seriousness of foodborne illness caused by astroviruses is dependant on the number of infective virus particles, moreover the age, genetic background, and general immune, health and nutrition status of the infected person, as well as the efficiency of sanitation systems. Thus foodborne diseases most seriously affect children, pregnant women, the elderly, and people already suffering from other diseases causing severe illness. Any food that has been handled manually and not at all or insufficiently heated is a possible source of astrovirus infection.115 Contamination of fresh produce by the ­astrovirus-containing fecal material frequently causes foodborne illnesses. The source of this contamination can vary from an infected person that has contact with food, to an entire harvesting area subjected to inefficiently or even untreated sewage or sludge (which is commonly used as fertilizer), and polluted water for irrigation.56 Bivalve molluscan shellfish can concentrate astroviruses from large volumes of water, allowing accumulation of virus from fecally contaminated water. 3.1.4.2 Antibody Acquisition and Prevalence Antibodies to astrovirus are generally acquired in early childhood. In 1978, an examination of 87 children under 10 years of age in the Oxford region of the UK demonstrated a rapid increase in antibody prevalence from 7% in 6–12-month-old babies, to 70% by school age.101 Astrovirus antibodies could be detected in 75% of the 10-year-old children. Young adults were also examined, and 77% had antibodies to astrovirus. In this study HAstV antibodies were detected by immunofluorescence of astrovirus infected cells; consequently, the prevalence of astrovirus in the population was likely underestimated. In a seroprevalence study among hospitalized children in the UK rates of 86% for serotype 1, 1% for type 2, 8% for serotype 3, and 6% for serotype 4 were reported.116

3.1.5 Laboratory Diagnosis Table 3.2 summarizes the current diagnostic methods for astroviruses. Most studies of astrovirus detection in food have focused on shellfish. Different protocols have been developed for the detection of astroviruses from foodstuffs; and comparative studies are needed to determine which assays should be recommended.117 The detection of astroviruses from any type of food sample without an effective concentration

39

Astroviruses

Table 3.2 The Current Diagnostic Techniques for Astroviruses Method Electron microscopy (EM)

Specimen

Sensitivity

Reagents

Negative stained fecal 106 to 107 virus specimens particles per gram of feces119 105 to 106 particles per gram of stool

• Organic solvents, resin, • Aqueous (or alcoholic) solutions of heavy metal stains

Enzyme immunoassay (EIA)

Clinical samples25

Molecular techniques (molecular probes, reverse transcriptionpolymerase chain reaction/RT-PCR/)

Both clinical and environmental samples69,118,120–125

• A group reactive monoclonal antibody (8E7) • Viral antigen • A polyclonal antiserum as the detector antibody • Commercial kit is available (DAKO Corporation,Carpenteria, CA) • Probe (fragment of astrovirus genome) in the case of molecular probes • Extracted astrovirus nucleic acid • Primers (oligonucleitodes from different regions of the genome)29,53,126,127 • Deoxinucleotides • Reverse transcriptase • Polymerase

Cell culturing

Both clinical and environmental samples

Immunoelectron microscopy (IEM)

Comparable sensitivity (91%) and specificity (98%) to IEM (105 to 106 particles per gram of stool)

• Molecular probes: 105 to 106 particles per gram of stool119 • RT-PCR: ten to 100 particles per gram of feces119

Media, trypsin

Limitations/Requirements • A  lack of specific antibodies to every serotype119 • Experienced microscopist • Only 10% of the astroviral particles in a given specimen display the distinctive surface star-like structure54 • Cannot be used for looking for the lower concentration of virus particles present in contaminated food or water

• Knowledge of human astrovirus genomes • The high variability in the genomic sequence does not allow the use of universal primers for all members of the family, Astroviridae36 • Fail to distinguish between infectious and noninfectious virus particles • Diagnostic methods for the detection of astroviruses or viral RNA in food and water have not been adopted to routine laboratories in most parts of the world • Naturally occurring inhibitors can hamper the PCR reaction. Therefore, the incorporation of an internal nucleic acid standard into each RT-PCR tube is important to identify inhibitors and eliminate false negatives • Requires the extraction of the viral RNA from the sample (e.g. with borate buffer, glycine solution, saline beef, proteinase K digestion, washing of food samples with guanidinium thiocyanate, adding PBS to the food sample, then extract with Freon) • More sensitive real-time RT-PCR has been developed • Have also been used to confirm the presence of astrovirus in EIA-positive samples28,95,104. Good correlation between EIA and RT-PCR has been observed, although it was demonstrated that in some cases RT-PCR has a higher sensitivity than EIA in detecting astrovirus28 • Have also been used to detect astrovirus genome in animals, such as minks128 and turkeys129 • Time-consuming, unreliable, expensive • Requires trypsin for isolation from primary specimens • Polluted water and shellfish samples are usually toxic for cell cultures

40

procedure and appropriate virus detection method is difficult and frequently unsuccessful. Several concentration methods have been developed in recent years. Concentration and purification of virions from shellfish rely on physicochemical procedures,118 such as polyethylene glycol (PEG) 6000 or 8000 precipitation, organic flocculation, or application of positively-charged virosorb filters. In addition, many different viral detection techniques have been used, ranging from tissue culture methods to nucleic acid hybridization. It is important to apply a method, which is flexible, inexpensive and can be used without extensive pH treatment or use of multiple reagents. Furthermore, it can be used in a minimally equipped laboratory and by staff with a minimal level of training. The traditional technique for detection of astroviruses is to isolate them in tissue culture. The isolation of astroviruses from food samples is not an easy task. Bacterial contaminations from shellfish also create difficulties in virus propagation in cell cultures. Treating samples with large amounts of antibiotics or chloroform might be effective, but toxic effects cannot be excluded. Therefore, at present, molecular techniques (such RT-PCR, real-time PCR, or immunomagnetic beads-PCR) offer the best alternative to develop sensitive and specific methods for the detection of astroviruses from environmental samples.130 Unfortunately, molecular techniques fail to distinguish between infectious and noninfectious virus particles, which may be a critical point in environmental virology and infection control.131 Although, the RNA genome does not remain intact without the protective capsid in an RNase-rich environment; therefore, if the genomic RNA could be detected by amplification, there is a good reason to suppose that viruses are infective. Virus detection in other foods, such as lettuce, strawberry, deli meats (ham, turkey, and roast beef), and green onion is not general.132

3.1.6 Treatment and Prevention Astrovirus gastroenteritis is generally characterized as a mild, self-limiting diarrhea with or without nausea and vomiting. The infection can strike down the individual for a few days, but does not require specific therapy. Although, more severe disease can develop in patients who have other medical problems, such as underlying gastrointestinal disease, malnutrition, immunodeficiency, coinfection with other pathogens(s) or prolonged illness. In addition, dehydration in young children may demand oral or intravenous fluid resuscitation. Intravenous immunoglobulin may be a beneficial adjunct in patients with severe immunodeficiency who have no response to conservative therapies,93 however, subsequent studies are required to determine effectiveness and to ascertain indications. Determinant of prevention of astrovirus infection is to break transmission, particularly in hospitals and other institutions, day-care centers, and families where person-to-person transmission is probable. Immunization against astroviruses is the ideal solution to prevent outbreaks, considering that

Molecular Detection of Foodborne Pathogens

astrovirus is a medically important pathogen.26,83 Vaccine development is not current, because the actual importance of astroviruses, as well as key factors of immunity are not well understood. Nevertheless, a prophylactic vaccine can play a remarkable role in reducing the incidence of food- and waterborne infections caused by astrovirus. Until then, general hygienic procedures, such as hand washing, disinfection of probably contaminated areas, surfaces (associated with feces or vomits containing astrovirus) with chlorine-based detergents, wearing of gloves for all points in the food chain where foodstuffs are handled manually, and appropriate treatment of potable water can prevent foodborne outbreaks and diseases due to astroviruses. It is also essential to not grow or wash foodstuffs in fecally contaminated water. Moreover, pasteurization procedures, disease control in animals, better regulation of shellfish beads, improvement in the safety of animals own feed and water supply, and protection from manure can prevent outbreaks; primarily because general hygienic procedures are not always sufficient to reduce astroviral infections and contamination.

3.2  Methods 3.2.1 Sample Preparation Frequently used virus concentration methods are summarized in Table 3.3, and general sample preparation and detection procedures of astroviruses from food samples are shown in Table 3.4. Food samples are usually obtained from cafeterias, coastal waters, as well as shellfish growing areas, and stored at 4°C, for a maximum of 24 hours until further processing. Generally, cold storage temperatures (2–8°C) delay senescence, product browning and growth of microorganisms in minimally processed fruits as well as vegetables, however promote the survival of astroviruses. Although, most studies of astrovirus detection in food have focused on shellfish. Shellfish are generally shucked, and the stomach and digestive verticula are removed by dissection, and frozen (–20°C) until analysis. For analysis, 20–25 g tissues need to be thawed on ice, then homogenized (e.g. in tryptose phosphate broth (TPB)-glycine buffer) and—in some cases—sonicated before virus concentration and nucleic acid extraction. In order to prevent cross-contamination, it is recommended to sterilize the homogenizing part of the blender with ethanol (70%) and to heat for 1 min after each sample. Some detailed sample preparation procedures are introduced in Section 3.2.3. Here we provide three step-wise protocols (Figures 3.3 through 3.5) for preparation of mussel, food and shellfish for RT-PCR detection of astroviruses.

3.2.3  Detection Procedures (i) Extraction of viral RNA: Viral RNA is prepared with one of the methods outlined in Figures 3.3 through 3.5. (ii) RT-PCR: The type-common primer pair Mon2/ PRBEG (Mon2, 5′ GCT TCT GAT TAA ATC AAT

41

Astroviruses

Table 3.3 Frequently used Virus Concentration Techniques from Foods Method

Reagents and Equipment

Underlying Priniciples

Hydroextraction

Homogenized sample in dialysis bag surrounded by polyethylene glycol

Viruses remain inside the bag, at 4°C, complete within 18–24 hours

Ultracentrifugation

Homogenized sample

Ultrafiltration

Homogenized sample, membrane

Adsorption to membrane filters Precipitation

Homogenized sample, positive/ negative filter Homogenized sample, PEG

Centrifuge at speeds above 50000  rpm Sample is driven through a membrane by applying pressure, viruses and macromolecules are retained Membrane filters retain viruses Add 8% PEG

Advantages/Disadvantages –Low recovery –Viruses are sometimes adsorbed to the dialysis membrane, –Toxic substances present in food may inactivate viruses during the 24-hour incubation period –Ultracentrifuge is very expensive –Good recovery, –Clogging of the filter

–Clogging, –Membrane coating components –Simple, cheap, –Good recovery

Table 3.4 General Sample Preparation and Detection Procedures of Astroviruses from Food Samples Concentration Homogenization, centrifugation Homogenization, sonification, centrifugation Homogenization in 1:7 (wt/vol) 10% TPB-0.05 M glycine (pH 9), centrifugation Homogenization, ultracentrifugation



Viral extraction or elution PEG 6000 centrifugation Freon TF centrifugation

Viral nucleic acid extraction Proteinase K, phenol-chloroform extraction, ethanol precipitation Mix of glass powder matrix and guanidine isothiocyanate

Detection

Reference

RT-PCR, hybridization

124

Nested RT-PCR

133

Chloroform extraction, PEG 6000 precipitation

Proteinase K digestion , phenolchloroform extraction, ethanol precipitation

RT-PCR, cell culture

33

Glycine buffer, PBS, centrifugation,

Guanidinium thiocyanate-silica particles

(nested) RT-PCR

134

TTT 3′; PRBEG, 5′ ACC GTG TAA CCC TCC TCT C 3′) targeting the hypervariable 3′ end of ORF2 region of astrovirus types 1–3 to 5–8, as well as Mon2/JWT4 (JWT4, 5′ GCA GAG AGC TTG TTA TTA AC 3′) for HAstV-4 are used for detection of HAstV by RT-PCR. These primers yield amplicons from 296 to 332 bp depending on the different HAstV types.136 (1) Prepare reverse transcription mixture (25 μl per tube) containing 0.1% bovine serum albumin, 0.4 mM dNTP mix (Promega, Madison, WI), 2 U AMV-RT (avian myeloblastosis virus RT, Promega, Madison, WI), 4 U RNasine (RNase inhibitor, Promega, Madison, WI), 0.05 μM dithiothreitol (Promega, Madison, WI), 2 μM





negative-strand primer (Mon2) and 5 μl RNA suspension in the reaction buffer (10 mM Tris, 50 mM KCl, 3 mM MgCl2, pH 8.3). (2) Incubate the tubes for 1 hour at 42°C. (3) Prepare PCR mixture (25 μl per tube) containing 10% dimethyl sulfoxide (Sigma, St. Louis, MO), Taq polymerase and 1 μM positive-strand primer (PRBEG, JWT4 or DM4 [DM4, 5′ CTA CAG TTC ACT CAA ATG AA 3′]) in the reaction buffer (same as the RT). (4) Transfer the total volume (25 μl) of cDNA from step 2 into each tube containing 25 μl PCR mixture. Perform PCR amplification with an initial denaturation at 94°C for 3 min, followed by 40 cycles of amplification; denaturation at 94°C for

42

Molecular Detection of Foodborne Pathogens

Homogenize (1 min, 9,500 rpm), then stir on magnetic stirrer (15 min) Sonicate (1 min, 100 W), centrifuge (10,000 × g, 90 min, 4°C)

Virus extraction

Grind the mussel tissue (3 min, 10,000 rpm), then add 100 ml 1 M borate-3% beef extract buffer pH 9

Adjust the pH to 7.3, then supplement the extract with 10% (final concentration) PEG 6000

Centrifuge (10,000 × g, 90 min, 4°C) Resuspend the pellet in 12 ml Na2HPO4 (pH 9) with vigorous magnetic stirring

Concentration

Incubate overnight at 4°C

Centrifuge (1,500 × g, 20 min, 4°C), then adjust the pH to 7.2 Extract RNA (RNA-PLUS purification kit; Bioprobe Systems, Montreuil, France or phenolchloroform extraction), resuspend the RNA pellet in 25 µl diethyl pyrocarbonate-treated water RT-PCR

Figure 3.3  Protocol I: preparation of mussel sample for RT-PCR detection of astroviruses (based on Traore et al., Appl. Environ. Microbiol., 64, 3118, 1998.) Wash the food (20–40 g) 3× with 40 ml PBS, mix for 5 min Freon extract (add 70 ml Freon, for 5 min, centrifugation [5,000 × g, 10 min, 4°C]) PEG precipitate (add 10% PEG 6000 and 0.3 M NaCl, incubate: 2 h, 4°C; centrifugation [7000 × g, 30 min, 4°C]) Extract the pellet with 8 ml TRIzol Clarification (centrifugation [8000 x g, 20 min, 4°C] Phenol/chlorophorm extract (add 1.6 ml chloroform to the aqueous phase, mix for 15 sec; incubation: 3 min, room temperature; centrifugation: 8000 × g, 20 min, 4°C; mix the aqueous layer with 4 ml isopropanol for 30 sec; incubation: 10 min, room temperature; centrifugation: 8000 × g, 20 min, 4°C) Precipitate viral RNA (wash the pellet with 8 ml 70 % ethanol; centrifugation: 7000 × g, 5 min, 4°C; air dry the pellet; suspend in 100 µl RNase-free water; store at –80°C RT-PCR amplification

Figure 3.4  Protocol II: preparation of food sample for RT-PCR detection of astroviruses (according to Schwab et al., Appl. Environ. Microbiol., 66, 213, 2000.) Homogenize 25 g shellfish in TPB-0.05 M glycine (pH 9.0–9.5) (5 min), then sonicate (2 min, 100 W) Concentration: centrifugation (3000 × g, 30 min, 4°C), adjust supernatants pH to 7.2–7.4 add PEG 6000 (8%;wt/vol), stir: 2 h, 4°C, then repeat centrifugation Detoxification: resuspend pellet in 5 ml 0.15 M Na2HPO4 (pH 9.0–9.5), sonicate 2 times (30 sec, 100 W) centrifugation (10000 × g, 30 min, 4°C), adjust supernatants pH to 7.4 Purification: add PBS to 15 ml final volume, then extract 5 times with Freon TF centrifugation (3000 × g, overnight, 4°C) over membrane filter Nucleic acid extraction: lyse with 6.6 M guanidine isothiocyanate, bind nucleic acid to silica particles, wash with guanidine isothiocyanate, ethanol and acetone Elution: TE buffer at 56°C, ethanol precipitate: add 0.1 vol. sodium acetate and 2 volume ethanol, leave at –70°C, 30 min, centrifuge at 20000 × g, 20 min, 4°C RT-PCR assay

Figure 3.5  Protocol III: preparation of shellfish for RT-PCR detection of astroviruses (based on Lees et al., Appl. Environ. Microbiol., 60, 2999, 1994.)

Astroviruses

1 min, annealing at 50°C for 1 min, and elongation at 72°C for 1 min. The final extension is at 72°C for 10 min. (5) Separate PCR products by agarose gel (3%) electrophoresis in Tris–boric acid EDTA buffer, pH 8.0, containing ethidium bromide (0.5 μg/ml), and visualize amplicons by UV-transillumination at 320 nm. (iii) Sequencing and phylogenetic analysis: The amplicons from the 3′ end of ORF2 obtained above are cloned into pGEM-T vector (Promega, Madison, WI) following the manufacturer’s instructions. Two different clones of the same RT-PCR amplicon are sequenced using fluorescein-labeled primers and commercial sequencing kit (SequiTerm EXCEL II Long-Read DNA Sequencing Kit-ALF, Epicentre Technologies, Madison, WI) on an automated sequencer (ABI 310, Applied Biosystems, Foster City, CA) following the manufacturer’s recommendations. Basic sequence manipulation and verification are performed using OMIGA software (v.2.0 Accelrys Co., San Diego, CA). Nucleotide sequences of the Hungarian strains are compared to available reference strains and a genotype is assigned based upon similarity scores. Viruses with 97–100% nucleotide identity in the 3’ end of ORF2 region of the genome are considered the same strain. In order to obtain a more accurate and reliable comparison, phylogenetic analysis is performed on representative strains using a longer, approximately 1.2-kb region (Mon2/DM4) of the 3’ end of ORF2. ClustalW v1.7 (http://evolution.genetics.washington.edu) is used to create multiple alignments of the amino acid sequences of the selected partial capsid sequences. The nucleotide sequences are added and aligned by GeneDoc v2.3 (http://www.psc. edu/biomed/genedoc) using the corresponding amino acid sequences as template, resulting in a consensus length of 1183 nucleotides terminating at the 3’ end of ORF2. A phylogenetic tree is constructed from the nucleotide sequence alignment using the maximum-likelihood algorithm in the program DNAML of PHYLIP v3.52c (http://evolution.genetics.washington.edu/phylip) running in a UNIX environment. The global rearrangement option is invoked and the order of the sequence input is randomized 50 times. The analysis is performed unrooted.

3.3 Conclusions and Future Perspectives Since astrovirus is one of the main four viruses that cause foodborne infections, studies are needed to estimate the burden and cost of illness caused by foodborne astroviral infections, especially among susceptible individuals. Therefore, better surveillance systems are required. Molecular investigation of astroviruses throughout the whole food chain and through populations is also essential. The burden of

43

foodborne illnesses due to astrovirus is highest in the elderly, and as a result of aging populations it will probably increase in the following years.137 The extreme stability of astroviruses in the environment and their highly infectious nature contribute to the ease of foodborne transmission. Infected foodhandlers play a well known role in transmission of astrovirus. Nevertheless, virus contamination may occur anywhere in the food chain, consequently the role of the infected agricultural laborer should be taken into consideration. The use of sludge as fertilizer and wastewater for irrigation increase the risk for viral contamination. There is a need to develop quick and simple molecular methods for detection of astroviruses from foodstuffs, and to urge rapid exchange of typing information between food laboratories and countries. New methods should be comparable and need to be standardized. Further studies are needed to evaluate the applicability of these methods in food microbiology.137 Molecular methods may play an important role to better understand foodborne astroviral outbreaks. Further studies are needed both to evaluate the mechanism of emergence of epidemic strains and the plausible link with animal astrovirus infections. There is a need for international cooperation as well as a well-organized public health system with the involvement of researchers to find solutions for foodborne astroviral outbreaks. Development of rapid detection methods to combine epidemiological and virological information is also necessary in the limitation of foodborne outbreaks. Contaminated products usually have a normal look, smell and taste, which present difficulties in the identification of astrovirus. Also, appropriate astrovirus detection methods are in general, not routinely available in food microbiology laboratories. While routine cell culture methods isolate astroviruses easily from blood or other sterile sites, specific methods are necessary to identify them from foodstuffs. A primary obstacle of linking epidemic outbreaks with foods is to detect the small amount of viruses that may be present in foodstuffs. In addition, by the time the infection is clinically manifest, the food under discussion has been consumed or discarded. In most countries the surveillance infrastructure for foodborne diseases of microbiological (or chemical) etiology, is weak or nonexistent. Moreover, in many countries, principally in the developing world, laboratory resources and skills to identify astroviruses are inadequate, and etiology-specific surveillance is often not possible. Even in the developed countries, laboratory-based surveillance is not well advanced. Absence of a simple and reliable diagnostic test to detect astroviruses in food makes surveillance difficult. Currently available routine monitoring systems focus on bacterial pathogens. Nowadays, directives establish no specific microbiological criteria concerning the presence of enteric viruses in food or water; nonetheless it has clearly been shown, that no correlation between the presence of HAstVs and indicators of fecal pollution was found. Further investigations are needed to establish possible correlations between fecal contamination and HAstVs as viral

44

contaminants.138 Nevertheless, the shortcomings of standard indicator organisms: fecal coliform bacteria, Escherichia coli, and Salmonellae as generally accepted indicators of fecal contamination of food, have been highlighted in several studies,139,140 and this has led to calls for a reassessment of quality guidelines based on these indicator organisms. The food industry and the scientific community should work together to develop an integrated plan of action to address foodborne astroviral infections.

References







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47 120. Chapron, C.D., Ballester, N.A., Fontaine, J.H., Frades, C.N., Margolin, A.B. Detection of astroviruses, enteroviruses, and adenovirus types 40 and 41 in surface waters collected and evaluated by the information collection rule and an integrated cell culture-nested PCR procedure. Appl. Environ. Microbiol. 66, 2520, 2000. 121. Elamri, D.E., Aouni, M., Parnaudeau, S., Le Guyader, F.S. Detection of human enteric viruses in shellfish collected in Tunisia. Lett. App. Microbiol. 43, 399, 2006. 122. Gabrieli, R., Macaluso, A., Lanni, L., Di Giamberardino, F., Cencioni, B., Petrinca, A.R., Divizia, M. Enteric viruses in molluscan shellfish. New Microbiol. 30, 471, 2007. 123. Grimm, A.C., Cashdollar, J.L., Williams, F.P., Fout, G.S. Development of an astrovirus RT-PCR detection assay for use with conventional real-time, and integrated cell ­culture/ RT-PCR. Can. J. Microbiol. 50, 269, 2004. 124. Le Guyader, F., Haugarreau, L., Miossec, L., Dubois, E. and Pommepuy, M. Three-year study to assess human enteric viruses in shellfish. Appl. Environ. Microbiol. 66, 3241, 2000. 125. Pintó, R.M., Villena, C., Le Guyader, F., Guix, S., Caballero, S., Pommepuy, M., Bosch, A. Astrovirus detection in wastewater samples. Water Sci. Technol. 43, 73, 2001. 126. Noel, J.S., Lee, T.W., Kurtz, J.B., Glass, R.I., Monroe, S.S. Typing of human astroviruses from clinical isolates by enzyme immunoassay and nucleotide sequencing. J. Clin. Microbiol. 33, 797, 1995. 127. Walter, J.E., Mitchell, D.K., Guerrero, M.L., Berke, T., Matson, D.O., Monroe, S.S., Pickering, L.K., Ruiz-Palacios, G. Molecular epidemiology of human astrovirus diarrhea among children from a periurban community of Mexico City. J. Infect. Dis. 183, 681, 2001. 128. Mittelholzer, C., Englund, L., Hedlund, K.O., Dietz, H.H., Svensson, L. Detection and sequence analysis of Danish and Swedish strains of mink astrovirus. J. Clin. Microbiol. 41, 5192, 2003. 129. Koci, M.D., Seal, B.S., Schultz-Cherry, S. Development of an RT-PCR diagnostic test for avian astrovirus. J. Virol. Methods 90, 79, 2000. 130. Tsai, Y-L., Tran, B., Sangermano, L.R. and Palmer, C.J. Detection of poliovirus, hepatitis A virus, and rotavirus from sewage and ocean water by triplex reverse transcriptase PCR. Appl. Environ. Microbiol. 60, 2400, 1994. 131. Abad, F. X., Pintó, R. M., Villena, C., Gajardo, R. and Bosch, A. Astrovirus survival in drinking water. Appl. Environ. Microbiol. 63, 3119, 1997. 132. Schwab, K.J, Neill, F.H., Fankhauser, R.L., Daniels, N.A, Monroe, S.S., Bergmire-Sweat, D.A., Estes, M.K., Atmar, L.R. Development of methods to detect “norwalk-like viruses” (NLVs) and hepatitis A virus in delicatessen foods: application to a food-borne NLV outbreak. Appl. Environ. Microbiol. 66, 213, 2000. 133. Green, J., Henshilwood, K., Gallimore, C.I., Brown, D.W.G., Lees, D.N. A nested reverse transcriptase PCR assay for detection of small round-structured viruses in environmentally contaminated molluscan shellfish. Appl. Environ. Microbiol. 64, 858, 1998. 134. Pina, S., Puig, M., Lucena, F., Jofre, J., Girones, R. Viral pollution in the environment and in shellfish: human adenovirus detection by PCR as an index of human viruses. Appl. Environ. Microbiol. 64, 3376, 1998. 135. Lees, D.N., Henshilwood, K., Doré, W.J. Development of a method for detection of enteroviruses in shellfish by PCR with poliovirus as a model. Appl. Environ. Microbiol. 60, 2999, 1994.

48 136. Meleg, E., Jakab, F., Kocsis, B., Bányai, K., Melegh, B., Szücs, Gy. Human astroviruses in raw sewage samples in Hungary. J. Appl. Microbiol. 101, 1123, 2006. 137. Koopmans, M., von Bonsdorff, C-H., Vinjé, J., de Medici, D., Monroe, S. Foodborne viruses. FEMS Microbiol. Rev. 26, 187, 2002. 138. Macaluso, A., Gabieli, R., Lanni, L., Saccares, S., Pana, A., Divizia, M. Enteric viruses and bacteriological parameters in molluscs. Ann Ig. 16(1–2), 237, 2004.

Molecular Detection of Foodborne Pathogens 139. Croci, L., De Medici, D., Scalfaro, C., Fiore, A., Divizia, M., Donia, D., Cosentino, A.M., Moretti, P., Costantini, G. Determination of enterovirus, hepatitis A virus; bacteriophages and E. coli in Adriatic sea mussels. J. Appl. Microbiol. 88, 293, 2000. 140. Le Guyader, F., Apaire-Marchais, V., Brillet, J., Billaudel, S. Use of genomic probes to detect hepatitis A virus and enterovirus RNA in wild shellfish and relationships of viral contamination and bacterial contamination. Appl. Environ. Microbiol. 59, 3963, 1993.

4 Avian Influenza Virus Giovanni Cattoli and Isabella Monne

Istituto Zooprofilattico Sperimentale delle Venezie

Contents 4.1 Introduction......................................................................................................................................................................... 49 4.1.1 Classification of AI Virus........................................................................................................................................ 49 4.1.2 Biology and Pathogenesis........................................................................................................................................ 50 4.1.3 Medical Importance and Zoonotic Implication of AI Viruses................................................................................ 50 4.1.4 Diagnosis of AI Viruses........................................................................................................................................... 51 4.1.4.1 Virus and Antigen Detection..................................................................................................................... 51 4.1.4.2 Serology for AI Diagnosis......................................................................................................................... 52 4.1.4.3 Molecular Tests......................................................................................................................................... 52 4.2 Methods............................................................................................................................................................................... 55 4.2.1 Sample Collection and Handling............................................................................................................................. 55 4.2.1.1 Selection of Samples to be Collected........................................................................................................ 55 4.2.1.2 Transportation and Storage of Specimens................................................................................................. 55 4.2.1.3 Handling of Specimens............................................................................................................................. 56 4.2.2 Samples Preparation for PCR Testing..................................................................................................................... 56 4.2.3 Detection Procedures............................................................................................................................................... 56 4.2.3.1 One Step RT-PCR for the Detection of Type A Influenza Viruses........................................................... 56 4.2.3.2 One Step RT-PCR for the Detection of AI Viruses Belonging to the H5 Subtype................................... 57 4.2.3.3 Two Step RT-PCR for the Subtype Specific Detection of H7 AI Virus.................................................... 57 4.2.3.4 Detection of Type A Influenza Virus by Qualitative Real Time PCR (M gene)...................................... 58 4.2.3.5 Detection of Type A Influenza Viruses of H5 HA Subtype by Qualitative One Step Real Time RT-PCR..................................................................................................................................................... 58 4.2.3.6 Detection of Type A Influenza Viruses of H7 HA Subtype by Qualitative Real Time PCR......................................................................................................................................... 59 4.3 Conclusions and Future Perspectives.................................................................................................................................. 59 References.................................................................................................................................................................................... 60

4.1  Introduction Recent outbreaks of avian influenza (AI) in birds occurring in Europe, in the Americas, Asia, and Africa, have provided field evidence of how challenging could it be to control this infection, particularly in densely populated poultry areas or in areas where free-range rural village poultry and backyard flocks are present.1–3 The early detection of AI in domestic and wild bird populations as well as in poultry ­commodities has been recognized as crucial to the implementation of timely and adequate prevention and control strategies. In the last decade, an increasing number of novel molecular technologies have become available to aid diagnosis of infectious diseases of animals and many of these have been applied to improve and accelerate the diagnosis of AI. In addition, more traditional diagnostic protocols, mainly based on classical viral culture methods and

i­ mmuno-enzymatic assays, have been revisited or improved in the recent years. The increased availability of diagnostic tests in conjunction with improved knowledge on the epidemiology and the pathogenesis of AI, is resulting in a modified approach to surveillance and diagnosis of this infection, with a wider application of the molecular-based technologies.4

4.1.1 Classification of AI Virus Influenza viruses have segmented, negative sense, single strand RNA genomes. The viruses are classified in the family Orthomyxoviridae. At present the Orthomyxoviridae family consists of five genera, only viruses of the Influenzavirus A genus are known to infect birds. Influenza A viruses are subdivided into subtypes based on the antigenic relationships in the surface glycoproteins, hemagglutinin (HA), and neuraminidase (NA). At present, 16 HA subtypes have been recognized

49

50

(H1–H16) and nine NA subtypes (N1–N9).5 Each virus has one type of HA and one type of NA antigen, apparently in any combination.

4.1.2  Biology and Pathogenesis Type A AI viruses are very widespread in nature; up to now they have been detected in more than 105 different species of wild birds from 26 families.6 Wild aquatic birds are natural reservoirs of these viruses and they can become infected by viruses of all HA and NA subtypes without showing signs of disease. In domesticated poultry AI viruses can be grouped in two pathotypes based on the clinical signs they may cause mainly in gallinaceous species: low pathogenic AI (LPAI) viruses and highly pathogenic AI (HPAI) viruses. LPAI viruses mostly cause infections of the respiratory and the enteric tract and the infection is subclinical in most avian species. Infections may be characterized by mild respiratory signs, some depression and reduced egg production. HPAI viruses not only replicate in the respiratory and enteric tract but also in endothelial cells throughout the body with spillover to adjacent parenchymal cells. Disease signs involve the respiratory and enteric tracts and many other organ systems, such as the central nervous system. Lesions are characterized by multiple haemorrhages in visceral organs and the skin, and mortality rates approach 100%. Infection with HPAI viruses thus leads to wide dissemination in the body and virus presence in many organs and edible tissues.7 To date, only viruses of H5 and H7 subtype have been shown to cause HPAI in susceptible species, but not all H5 and H7 viruses can be classified as HPAI. It appears that H5 and H7 HPAI viruses arise by mutation after an AI virus precursor of low pathogenicity (LPAI) has been introduced into poultry. It follows that all HPAI viruses should have a LPAI progenitor, although the latter have only been identified in a limited number of cases.8 For all influenza A viruses the HA glycoprotein is produced as a precursor, HA0, which requires post translational cleavage by host proteases before it is functional and virus particles are infectious.9 The HA0 precursor proteins of AI viruses of low virulence for poultry (LPAI viruses) have a single arginine at the cleavage site and another basic amino acid at position -3 or -4 from the cleavage site. These viruses are limited to cleavage by extracellular host proteases such as trypsin-like enzymes and thus restricted to replication at sites in the host where such enzymes are found, i.e., the respiratory and intestinal tracts. HPAI viruses possess multiple basic amino acids (arginine and lysine) at their HA0 cleavage sites, either as a result of apparent insertion or apparent substitution, and appear to be cleavable by an intracellular ubiquitous proteases. HPAI viruses are able to replicate throughout the bird, damaging vital organs and tissues, which results in disease and death.10,11 The factors that bring about mutation from LPAI to HPAI are not known. In some instances mutation seems to have taken place rapidly (at the primary site) after introduction in

Molecular Detection of Foodborne Pathogens

poultry possibly through the wild birds,12,13 in others the LPAI virus has circulated in poultry for months before mutating.14 Therefore, it is impossible to predict if and when this mutation will occur. However, it can be reasonably assumed that the wider the circulation of LPAI in poultry, the higher the chance that mutation to HPAI will occur. HPAI viruses are not necessarily virulent for all species of birds and the severity of the clinical signs may vary with bird species, age of the host and virus strain.15,16 In particular, ducks rarely show clinical signs as a result of HPAI infections although there are reports that some of the Asian H5N1 viruses have caused disease and the HPAI viruses A/duck/Italy/2000 H7N1 and A/chicken/Germany/34 (H7N1) have been reported to cause disease and death in naturally and experimentally infected waterfowl.15,17 Bird to bird transmission of AI viruses is complex and it largely depends on the virus strain, host species and environmental factors.18 LPAI viruses are mainly excreted with feces, through the cloaca. Viral shedding through the respiratory tract is also considered important, at least for some species or some strains, as the Asian HPAI H5N1. For this virus, and similarly for other HPAI viruses, transmission from poultry to humans is supposed to occur primarily through direct contact with secretions of the upper respiratory tract, infected feces, feathers, organs, and blood of infected animals. Inhalation of contaminated dust or droplets can be an alternative transmission route.19

4.1.3 Medical Importance and Zoonotic Implication of AI Viruses Until recently, direct infection of humans with AI viruses had not been considered significant. Human cases were sporadically reported between 1959 and 1996 with only three documented cases, two in the USA (HPAI and LPAI H7N7) and one, likely to be of laboratory exposure-origin, in Australia (HPAI H7N7). In all cases the patients recovered and the main clinical sign was characterized by self-limiting conjunctivitis.3,8 However, starting from 1996, a series of events has raised the concerns on the zoonotic potential of AI infections. With some exceptions, since 1996 almost all the reported cases of AI virus infection in humans have been caused by HPAI viruses belonging to the H5 or H7 subtypes directly transmitted from infected birds to humans. Among the LPAI viruses, the first documented case of avian to human transmission was described in 1996 and caused by a LPAI H7N7 virus. The virus was isolated in England from the eye of a woman suffering from conjunctivitis. This person had ducks and the virus was shown to be genetically 100% of avian origin.20,21 In 1999, LPAI viruses belonging to the H9N2 subtype were isolated from two young girls in Hong Kong.22 They were suffering from an influenza-like syndrome and fortunately they recovered with no serious consequences. Subsequently, isolation of H9N2 viruses in human beings was reported in the Peoples Republic of China on five occasions during 1998.23 In 2003, LPAI H7N2 was isolated from a patient with respiratory symptoms in the US. Also in this case, the patient recovered.3

Avian Influenza Virus

In two other circumstances, evidence of contacts between LPAI viruses and humans were only detected serologically. A serological survey in human beings potentially at risk of exposure during the 2002–2003 LPAI H7N3 epidemic in Italy revealed the presence of specific antibodies in 3.8% of serum samples collected in poultry workers.24 Serological prevalence with regard to H9N2 was also revealed by the hemagglutination inhibition test in the human population at risk of exposure in Iran.25 However, this latter finding needs confirmation by means of other serological tests. Certainly, more significant and severe are the human infections caused by HPAI viruses. The first documented evidence on how serious could be the consequences of avian to human transmission of HPAI viruses occurred in 1997 in Hong Kong. In that year, the HPAI H5N1 circulating in the domestic poultry was capable of infecting 18 people, causing the death of six of them.26 Viruses belonging to the same antigenic subtype, H5N1, and genetically related to the 1997 viruses re-emerged in Hong Kong in 2003.27 In that year, this HPAI virus circulating in poultry in South East China began to spread westward among wild and domestic birds throughout Asia, reaching Europe and Africa in 2005 and 2006. Since then, the continued infections of humans and other mammals, such as felines, caused by this virus have caused great concern over the capabilities of H5N1 to cross the species barrier and to potentially become easily transmissible among humans. To date (2nd February 2009), a total of 404 confirmed cases of HPAI H5N1 infections in humans and 254 human deaths have been reported to the World Health Organization (WHO) (available at http://www.who.int/csr/disease/avian_influenza/ country/cases_table_2008_04_17/en/index.html). Other HPAI viruses belonging to the H7 subtype were reported as etiological agents of severe human infections to a minor extent compared to HPAI H5N1. During the 2003 outbreak caused by the H7N7 HPAI virus in poultry in The Netherlands, 82 cases were reported in humans. Generally, symptoms were described as influenza-like illness and/or conjunctivitis, but one fatality also occurred.28 In Canada, persons involved in outbreaks management suffered from conjunctivitis, headache, and flu-like syndrome. The H7N3 HPAI virus, responsible for the outbreak in poultry, was confirmed to be the causative agent of the disease. Fortunately, no fatal cases occurred.29 While such human infections generally result from direct and intensive contact with infected or diseased poultry, other routes of infection such as consumption of edible tissues from infected avians or contact with contaminated water have been suggested as possible sources of infection.

4.1.4  Diagnosis of AI Viruses Surveillance and monitoring programmes have been implemented in many countries around the world as a result of the global spread of AI viruses and of the subsequent implications on public perception and animal health issues. These programmes are mainly targeting the wild birds—considered as the main reservoirs—and the poultry population. The main aims of these

51

programmes are to detect and control AI viruses in the poultry compartment, thus preventing their spread to human beings. Therefore, nowadays laboratory testing is mainly applied to trace viral circulation in a given area or in a susceptible population in order to implement an early warning system, in addition to diagnose the presence of the virus in a diseased flock or animal or in poultry derived products. This implies the use of rapid, sensitive and, possibly, cost-effective laboratory tests adaptable to very high throughputs.4 4.1.4.1  Virus and Antigen Detection Traditionally, laboratory protocols for the detection and the identification of AI viruses were based on virus isolation (VI) in SPF eggs or in cell cultures. The application of these methods of laboratory investigation is mainly limited by the fact that they are not flexible to a sudden increase in demand, are not cost-effective and often require a long processing time. In fact, these methods are time-consuming and require a minimum of 12 days before a negative result may be issued. Office International des Epizooties (OIE) and EU official methods require two blind passages in eggs of 6 days, before the sample is considered negative.30,31 At present, there seems to be only limited space for improving the time-efficiency of these methods. What appears to be a major bottleneck is the obtainment of suitable substrates for VI. The primary cell cultures and the continuous cell lines tested so far provide variable results, mainly strain to strain dependant and, in general, they are less sensitive than SPF eggs. These are expensive and not always easily available. The use of eggs derived from AI and Newcastle disease virus (NDV) specific antibody negative (SAN) parent stocks are considered as an alternative method in the OIE manual30 and, more recently, in the EU manual.31 VI implies the replication in laboratory of viable viral particles to a significant concentration thus, biosafety, and biocontainment should be regarded as a priority for laboratories in which AI VI is performed. Despite these major difficulties, VI in fowl’s eggs still remains the gold standard for AI virus detection. Its sensitivity is equal or often superior to many alternative tests. In addition, genetic or antigenic variation of the viruses, as well as the presence of ­contaminants or PCR inhibitors in the samples, can impair the efficiency of molecular and immunoassays, but they have minor impact on VI. Under certain circumstances, it might be desirable to test a certain number of samples in a short period of time. In this case, antigen capture immunoassays can be ­considered a very useful diagnostic tool. They are very easy to use, do not require sophisticated or expensive equipment and, in many instances, they can be applied on-site, thus avoiding the time-consuming and delicate phase of sample preparation and shipment. In many cases, test results can be available within minutes. To date, most of the antigen capture tests available on the market target the type A influenza virus nucleoprotein (NP), thus detecting any type A influenza virus. With the exception of one H5 subtype specific test for veterinary use, this kind of

52

Molecular Detection of Foodborne Pathogens

assay does not provide any indication on the subtype involved. Furthermore, no indication on the pathotype (i.e., HPAI vs LPAI) can be obtained. Their main limitations consist in the unsatisfactory sensitivity compared to VI and molecular tests, and their unit cost.32,33 Due to their low sensitivity, sampling numbers should be increased before the presence of viral circulation in a given population can be ruled out. For the same reason, excessive dilution of the samples by pooling of swabs should be avoided. These features make this kind of assay unsuitable for monitoring and early detection programmes, where large number of samples must be cost-effectively screened with sensitive testing procedures. 4.1.4.2 Serology for AI Diagnosis Serology represents a relatively inexpensive and practical methodology to assess the circulation and the prevalence of influenza viruses in poultry. It does not provide direct evidence on the presence of the virus but, in the framework of a surveillance effort in domesticated birds, serological diagnosis is considered a suitable approach to monitor the AI-free status of a given region or farm. On the contrary, the application of the serological methods available today in wild bird surveillance may generate information of very limited use, particularly for HPAI surveillance.34 In fact, serology does not provide information concerning the pathotype and wild birds, specifically wild waterfowl, which are commonly infected by LPAI viruses, therefore serologically positive. During monitoring or surveillance programmes, large numbers of animals have to be tested in order to guarantee acceptable statistical significance. In recent times the agar gel immuno-diffusion (AGID) test has been superseded by commercial ELISA tests. Both systems detect antibodies to

the group antigen of influenza A viruses, and therefore are unable to give any indication on the virus subtype causing infection. These tests find their primary application in monitoring poultry flocks, although most of them lack validation data for minor species (ducks, geese, quails etc). In addition, AGID is not suitable for testing sera of waterfowl as the latter do not produce precipitating antibodies.35 The hemagglutination inhibition test is a simple, robust, and fully validated test that generates qualitative and ­quantitative information on antibodies that are a result of vaccination or of infection in most avian species. It is more labor-intensive than ELISA or AGID tests, but it yields ­information that under certain circumstances is more valuable and useful for managing field situations. 4.1.4.3  Molecular Tests In implementation of surveillance and monitoring programmes for AI, cost-effective testing procedures capable of facing high and constant workflows are necessary. During AI outbreaks the main problems encountered by a diagnostic laboratory are represented by a sudden increase in sample testing and an increased pressure for faster turn-around-time (TAT), both combined with high quality test performances and cost effectiveness.36 In this respect, the possibility of diagnosing AI by using molecular methods offers important advantages compared to other protocols, such as VI and ELISA. For this reason, in the recent past there has been a significant increase in the development and application of testing procedures for the detection of AI viral RNA. Several RT-PCR and real time PCR protocols have been published in scientific journals (see Table 4.1 for references) and the most recent methodologies such as NASBA,37,38 LAMP-PCR39,40 and pyrosequencing41,42

Table 4.1 Main Representative PCR-Based Protocols for the Detection of AI Viruses Published Since 2000 in International Scientific Journals Target End point RT-PCR Type A influenza virus Type A influenza virus, H5 and H7 subtypes Type A influenza virus, H5 and H7 subtypes Type A influenza virus and Newcastle Disease virus Type A influenza virus, avian pneumovirus and Newcastle Disease virus Type A influenza virus, H5, H7, H9 subtypes Type A influenza virus, H5, H7 and H9 H5 AI subtypes H7 AI subtypes H5 and H7 subtypes

Assay

One step RT-PCR Two step RT-PCR

Notes

Reference

82 83

One step RT-PCR-ELISA Two step duplex RT-PCR

Positive samples confirmed by dot-blot hybridisation. Emi-nested PCR for H7 subtype. Laboratory evaluation. Partially validated on Eurasian lineage. Validated assay.

One step multiplex RT-PCR

Laboratory evaluation.

85

One step type A RT-PCR and one step multiplex RT-PCR (H5, H7, H9) Two step, multiplex RT-PCR One step RT-PCR Two step RT-PCR One step multiplex RT-PCR

Laboratory evaluation.

86

Laboratory evaluation. Ring test evaluation on Eurasian strains. Ring test evaluation on Eurasian strains. Limited clinical validation.

87 56 56 88

44 84

(Continued)

53

Avian Influenza Virus

Table 4.1  (Continued) Target

Assay

Notes

H5 subtype of the H5N1 HPAI virus H1-H16 AI subtypes H1-H16 AI subtypes H1-H15

One step RT-PCR RT-PCR and sequencing RT-PCR and sequencing RT-PCR and sequencing

Limited clinical validation. Limited validation. Laboratory evaluation. Laboratory evaluation.

Real time RT-PCR (rRT-PCR) Type A influenza virus

One step rRT-PCR

Type A influenza virus Type A influenza virus

One step rRT-PCR One step rRT-PCR

Type A influenza virus

Two step rRT-PCR

Type A influenza virus, H5 and N1 Type A influenza virus, H5 and H9

One step multiplex rRT-PCR One step multiplex rRT-PCR

Type A influenza virus, H9 and N2

One step multiplex rRT-PCR

Type A and B influenza virus, H5

One step multiplex rRT-PCR

Type A and B influenza virus, H5 and N1

One step multiplex rRT-PCR

H5, H7, H9 AI subtypes

One step rRT-PCR

H5 and H7 subtypes

One step multiplex rRT-PCR

H5 AI subtypes

One step rRT-PCR

H5N1 HPAI virus of the Qinghai lineage

One step rRT-PCR

H5 subtype of the H5N1 HPAI virus

Two step rRT-PCR

H5 subtype of the H5N1 HPAI virus

Two step multiplex rRT-PCR

H5 subtype of the H5N1 HPAI virus

One step rRT-PCR

N1 subtype

One step rRT-PCR

Hydrolysis probe. Validated assay. MGB-hydrolysis probe. Laboratory evaluation. Light upon extension fluorogenic primers. Laboratory evaluation. SybrGreen chemistry. Limited field evaluation. MGB-hydrolysis probes. Laboratory evaluation. Hydrolysis probe. Laboratory evaluation. SybrGreen 1 dye. Validated assay Discrimination between type A and B not possible. Laboratory evaluation. Hydrolysis probes. Limited clinical validation on human specimens. Hydrolysis probes. Validated assay for Eurasian lineage. Hydrolysis probe. Validated assay for American lineage. Hydrolysis probes. Validated assay for Eurasian lineage. Hydrolysis probe. Validated assay. MGB-hydrolysis probes. Laboratory evaluation on one human specimen. Hydrolysis probes targeting two distinct regions of the HA molecule. Validated on human specimens of Hong Kong and Vietnam origin. Hydrolysis probe. Limited clinical validation on human specimens. MGB-hydrolysis probes. Validated assay on N1 subtypes pf the Eurasian lineage

have been applied, in many cases successfully, for the detection and typing of AI viruses. However, the use of these latter techniques is limited to research purposes at the moment. With reference to the application of nucleic acid amplification protocols, sample processing appears less cold-chain dependant, as the preservation of cellular integrity and virus viability is not essential for these assays. The possibility to detect AI viral RNA in samples containing inactivated viral particles due to prolonged storage or shipment, or in samples treated to eliminate viral infectivity increases the chances to diagnose the disease in specimens collected in remote area of the world and addresses the biosafety issues. Reagents are also available to better preserve the integrity of a fragile molecule as the RNA at environmental temperatures.43 Thus,

Reference 89 61 60 59

45, 56 90 91 92 93 94 95 96 97 46 45 54 57 98 99

42 100

unlike VI, molecular techniques can also be applied in small laboratories, providing that the basic equipment is available. This can contribute to the extension of a diagnostic laboratory network in the affected area and to the reduction of the TAT by avoiding the submission of the samples to a distant central and fully equipped laboratory of virology. In terms of analytical and diagnostic sensitivity and specificity, the data available concerning the molecular tests for AI can be considered as optimal for their application in large scale diagnosis during an outbreak, showing values of diagnostic sensitivity, and specificity close to or above 90%.32,44,45 An excellent agreement with the VI was reported during a LPAI outbreak investigation32 and in clinical validation of real time protocols.46 Data derived from experimental study32

54

indicate that RT-PCR based methods are capable of revealing viral RNA in tracheal swabs of experimentally infected birds for 10–15 days after challenge, similarly to VI. This period of time covers the first 7–10 days post-infection, when serology is not of diagnostic aid due to an undetectable immune response. Similarly, other studies provided experimental evidence of the usefulness of PCR-based techniques as an alternative to qualiquantitative VI methodologies.47,48 Molecular techniques can provide a variety of data useful for surveillance, monitoring, and outbreak investigation. Once the RNA is extracted, it is possible to gain information not only concerning the presence of virus in the clinical specimen, but also about the HA and NA gene segments, the pathotype (LPAI vs HPAI virus) and other genomic sequencing data that can be used for molecular epidemiology. Most importantly, expensive, and time consuming in vivo tests for pathogenicity can be avoided, preserving animal welfare. However, the recent and extensive applications of these kinds of molecular assays in the field of AI diagnosis have highlighted some drawbacks. The costs related to the equipment and reagents needed for PCR and real time PCR testing are still significant, although they decreased in recent times, mainly as a result of the widespread use of these technologies and the subsequent marketing competition. Nucleic acids amplification methodologies are generally extremely sensitive assays, making them prone to easily reveal cross contamination of samples, leading to false positive results. Mishandling of the extracted RNA, improper use of reagents or use of nonsterile, non RNAse-free disposables, or inadequate reference controls, may result in false negative test response. AI viruses exhibit a significant degree of genetic variability, particularly in certain important regions of the genome, as for segment 4 (HA) and 6 (NA). This might lead to diagnostic failures of some molecular tests based on primers and probes targeting these hyper variable regions when applied on mutated or new emerging viruses. A recent example for this is the PCR detection failure of viruses belonging to the H7 subtypes occurred during wild bird surveillance program in the US. In this case, a few nucleotides mismatches in the PCR probe targeting the HA molecule were the likely cause of failure.49 Consequently, the recent initiatives concerning AI sequence data sharing50 are crucial in the understanding of viral evolution and in updating probes and other molecular diagnostic tools as long as the viruses mutate. Considering the extreme sensitivity of many nucleic acid amplification assays, some samples tested positive by a given molecular test might not be confirmed by any other test applied on the same sample, including VI.51 Therefore, the adoption of fully validated protocols and harmonized test is mandatory. Examples of unexpected false positive results may indeed occur after extensive field application.52 Since AI is now threatening almost every continent, these considerations should seriously be taken into account when planning official guidelines and diagnostic protocols. Public institutions and private biotechnology companies are currently investing resources in research and development of

Molecular Detection of Foodborne Pathogens

more stabilized PCR reagents, robust molecular protocols and easy-to use, flock-side molecular tests.53 Currently, the most common types of molecular tests used for AI detection are RT-PCR or real time RT-PCR (rRT-PCR)based protocols (Table 4.1). Schematically, these can be further subdivided in protocols for the so-called “generic” detection of type A influenza viruses and protocols for the detection and identification of specific type A influenza virus subtypes. The common genomic targets of the first type of molecular tests are well conserved segments of the viral genome located in the genes encoding for the matrix proteins (M1 and 2) or the NP. Since these proteins are antigenically and genetically conserved regardless of the virus subtype, these types of tests are virtually capable of detecting type A influenza viruses belonging to subtypes H1–H16. Based on the available literature, these protocols exhibit high sensitivity and specificity, with higher performances of the rRT-PCR tests compared to the RT-PCR tests. For these reasons, these type A influenza tests are used as screening tests to evaluate the infectious status of an animal, flock or area (see Table 4.1 for references). Infections caused by AI viruses belonging to the H5 and H7 subtype are of major concern for public health and economic impact. Regardless of their pathogenicity, which may vary, they are notifiable diseases of animals, according to the current international legislation for animal health.30 Therefore, their occurrence must be detected promptly and immediately reported to the national and international veterinary authorities. As a consequence, all the samples tested positive in the screening for type A influenza virus should be PCR tested for the identification of these two subtypes and/or immediately submitted for VI attempts.31,36 Several protocols have been described for the specific PCR detection of H5 and H7 directly in clinical specimens (Table 4.1) and some of them properly validated.45,46,54 Phylogenetic studies55 demonstrated that H5 and H7 sequences could be divided into two major groups, related to the geographical origin of the viruses. Thus, so called “American” and “Eurasian” lineages were described among avian H5 and H7 viruses. These groups reflect the genetic variation observed in the targeted genes and this has also ­influence in the development and application of specific diagnostic assays. In fact, molecular tests designed on viruses belonging to the American lineage generally exhibit poor performances, in term of sensitivity, when applied on the Eurasian strains and vice versa.45,56 The major public health and veterinary concern raised by the spread of the Asian H5N1 HPAI also contributed to the development of several molecular tests specifically targeting this dangerous virus (Table 4.1). In many instances, these protocols are duplex RT-PCR or rRT-PCR tests targeting the H5 and the N1 gene segments of this specific virus, but sometimes they result in poor performances when applied on different H5 strains. One protocol has been described targeting one specific H5N1 HPAI genetic sublineage.57 LPAI viruses belonging to the H9 subtype are also of major interest for the economic losses they can cause to the

Avian Influenza Virus

poultry industries58 in addition to their sporadic capabilities to be transmitted to humans, fortunately without serious consequences.22 These reasons have also led to the development of rapid molecular tests for its detection in clinical specimens (see Table 4.1 for references). In some instances, there is the need to determine the specific subtype of type A PCR positive samples, once the involvement of the H5, H7, and H9 subtypes has been ruled out. RT-PCR protocols designed on conserved regions of the HA molecule were developed and subsequent sequence analysis of the PCR product should enable the identification of the H1–H16 subtype.59,60,61 These protocols are useful for rapid subtyping of viral isolates, but the lower sensitivity of these methods may represent a limit for their direct application on clinical specimens. Currently, none of the published molecular tests have been specifically validated for AI detection in poultry-derived products such as meat, feathers, etc. Recently, a protocol describing the RNA extraction procedure and the subsequent RT-PCR protocol for the detection of AI RNA from manure has been described.62

4.2  Methods 4.2.1 Sample Collection and Handling 4.2.1.1 Selection of Samples to be Collected Protocols concerning sample collection for AI are mainly focused on the detection of the virus in holdings suspected to be infected, thus, in living, moribund or recently dead birds. As a reference, the EU diagnostic manual for AI, adopted in 2006 by the EU31 or the OIE Manual for Diagnostic Tests and Vaccines for Terrestrial Animals30 provide concise but practical guidelines concerning sample collection and transportation. For molecular testing, viable viruses are not requested. However, the target RNA molecule is extremely fragile and its degradation due to improper sample handling and storage may result in false negative results. The targeted replication sites of LPAI viruses are the respiratory (mainly trachea and lungs) and the digestive (intestines) tract. HPAI viruses are disseminated throughout the whole body tissues. Both, LPAI and HPAI viruses, are shed via the upper respiratory tract and feces but the quantity of shed viral particles and the duration of shedding may vary according to host species, host immune status, and virus strains.63,64 Thus, tissue specimens from trachea, lungs, and intestines or oropharyngeal, tracheal, or cloacal swabs are suitable samples for AI detection in carcasses and/or living birds. In addition, for HPAI viruses, specimens from brain and other internal organs, such as spleen, heart and kidney, can be collected. Intestinal contents of fresh feces also may contain high viral loads.65–67 However, presence of potential PCR inhibitors, bacterial or fungal contaminations and proteases impose particular attention when dealing with this type of material as false PCR results (positives or negatives) may occur. In case of HPAI infections, the virus causes viremia and therefore can be present also in the blood of recently infected birds.68–70 Data on the duration and intensity of the

55

viremic period are scarce and these parameters can vary greatly according to the virus strain, its pathotype or the host involved. For example, virus titres in experimentally infected chickens varied from 101.4 EID50/ml for H5N2 HPAI69 to 108 EID50/ml for H5N1 HPAI.68 In another study, ducks experimentally inoculated with A/H5N1 HPAI revealed virus titers in blood ranging from 100.7 EID50/ml to 102.3 EID50/ml.71 For LPAI viruses, replication occurs at the epithelial surfaces of the respiratory and intestinal tracts; therefore viremia is not expected for this pathotype. However, in a few occasions H9 or H7 LPAI virus genomes have been detected in blood and viable viruses were isolated.72,73 Based on the above mentioned data, blood does not represent an ideal sample for AI detection but it can be considered as a potential source of environmental contamination during handling and processing of blood samples collected from suspected HPAI infected animals or during slaughtering of poultry in the pre-clinical phase of the disease. Viremia and viral replication in endothelial cells occurring in HPAI infections contribute to the colonization of muscles and other edible tissues (i.e., liver) in infected birds. Muscles and meat of domestic birds were found to be HPAI infected during field investigations on imported poultry products74 as well as experimental infections.69,75 Incomplete evisceration or contamination from lungs and intestine during slaughtering procedures may lead to LPAI or HPAI contaminated meat or carcasses, as recently demonstrated.76 Therefore, muscles (particularly breast and tight muscles) or respiratory and intestinal tissues still present in poultry carcasses represent suitable specimens for the detection of both HPAI and LPAI viruses. 4.2.1.2 Transportation and Storage of Specimens Swabs, tissue specimens, or feces should be immediately submitted to the laboratory for testing. Soon after their collection, samples should be refrigerated on ice or with frozen gel packs. In case a submission delay (>24 h) to the laboratory is expected, samples should be frozen in dry ice, liquid nitrogen or -80°C. Repeated cycles of freezing and thawing must be avoided to prevent reduction of the viral load, cell lysis, and consequent RNA degradation. Storage of allantoic fluids of eggs inoculated with H5 and H7 HPAI, and LPAI viruses in the guanidine-based lysis buffer included in two commercial RNA extraction kits preserved the suitability of the original RNA template for real time PCR amplification up to 7 days at + 4°C; ambient temperature and at + 37°C.77 Importantly, the same lysis buffers were able to inactivate both the HPAI and LPAI viruses tested after 4 h, thus increasing the biosafety of the handled specimens. In one other investigation,78 end point RT-PCR amplification was still possible after 2 weeks storage at ambient temperature of bird fecal samples spiked with influenza A viruses using guanidine buffer, commercial preservative or alcohols (ethanol or isopropanol) as preservatives. PCR products up to 521 bp were obtained in samples preserved with guanidine or commercial buffers and up to 206 bp in samples

56

preserved with alcohols. Obviously, all these procedures will make parallel VI attempts impossible. In case VI is necessary to confirm results of molecular testing or to assess viral infectivity, viral transport medium (VTM) should be used. VTM are generally based on phosphate saline buffered solutions (PBS, pH 7.0–7.4) or protein-based media, such as brain heart infusion (BHI) or tris-buffered tryptose bacteriological media supplemented with antibiotics and/or antifungals.31 VTM supplementation with glycerol (10–20%) contributes to better preserve sample stability and integrity, particularly during prolonged storage at low temperatures. 4.2.1.3 Handling of Specimens Probably only one documented case of human infection caused by an AI virus following laboratory exposure has been reported so far in the scientific literature.79 The case occurred in Australia and consisted of conjunctivitis caused by an H7N7 HPAI virus. In contrast, several cases of conjunctivitis, influenza-like illness and one fatal case of acute lung infection were reported during the H7N7 HPAI poultry outbreaks in 2003 in The Netherlands.28 People confirmed to be infected were somehow involved in the outbreak management. Adequate protective measures should therefore be adopted during collection and handling of samples suspected to contain AI viruses, at the farm level, at the slaughterhouse and within the laboratory. Personal protective items (PPI), such as lab-coats, goggles, disposable gloves, should be properly worn during necropsies and collection of samples from animals suspected to be AI-infected. Orthomyxoviruses are identified as biological agents of biohazard class 2.80,81 According to the WHO recommen­ dations,81 they should be manipulated in a BSL2 laboratory, adopting BSL3 work practices. Good ­laboratory practices should be applied during the whole process of ­sample testing at laboratory level. Useful and practical guidelines and comprehensive information on biosafety-related issues in the laboratory can be found in the EU manual for AI and on the WHO website.31,80,81

4.2.2 Samples Preparation for PCR Testing The following procedures for sample preparation can be followed prior to submitting the specimen for RNA extraction and subsequent RT-PCR amplification. Organs and tissues (brain, trachea, lungs, intestine): extract the RNA from tissue homogenate. Using sterile scissors or surgical blades, cut small blocks (approximately 2–5 mm × 2–5 mm) of tissues. If possible, tissues blocks should be frozen in liquid nitrogen to better preserve RNA integrity and to facilitate tissue disruption. Animal tissues can then be disrupted simply by sterile pestle and mortar. Fiber-rich tissues (e.g., lung and trachea) may require the addition of sterile quartz powder or sand to better disrupt tissue cells. To facilitate the disruption and the homogenization process, 300–500 µl of sterile PBS can be added. This will make also possible the preparation of aliquots for other types of test (i.e., VI),

Molecular Detection of Foodborne Pathogens

starting exactly from the same homogenate. Homogenization is carried out simply using a syringe and needle. Alternatively, a commercially available, automatic homogenizer can be used. Add the requested amount of this suspension to the lysis buffer of the RNA extraction kit according to the manufacturer’s instructions. Cloacal swabs, tracheal, or oro-pharyngeal swabs: dilute swabs in PBS (max 1 ml) and extract the RNA from this suspension. It is possible to pool the samples (up to ten tracheal swabs/pool or five cloacal swabs/pool). Vortex briefly. Add the requested amount of this suspension to the lysis buffer of the RNA extraction kit according to the manufacturer’s instructions. Feces: a suspension is prepared by adding one volume of feces to four volumes of sterile PBS. Add the requested amount of this suspension to the lysis buffer of the RNA extraction kit according to the manufacturer’s instructions. Allantoic fluid: simply add the requested amount of sample (allantoic fluid) to the lysis buffer of the RNA extraction kit according to the manufacturer’s instructions. RNA extraction: several methods for the manual or robotic extraction of the RNA exist. Many commercial kits are available, some of them developed and optimized for the extraction of the nucleic acids on specific matrixes, such as tissues, blood, and stool. However, to facilitate the organization of sample processing within the laboratory and make it more cost-effective and practical, only some kits are presented in this chapter, which can be used on different matrixes with satisfactory results. The kits listed should be considered as examples, representatives of the most common types used in different laboratories or described in several scientific journals. They have been evaluated in many laboratories working on AI. The use of other kits not included in the list is possible, providing their performances are methodically evaluated. Kits widely used for RNA extraction: NucleoSpin® RNA II (Macherey-Nagel GmbH & Co., Germany); RNeasy® MiniKit (Qiagen Gmbh, Germany); High pure RNA isolation kit (Roche Applied Science, Germany), not recommended for feces; MagMax (Ambion/Applied Biosystems), for swabs or other liquid matrix. Not recommended for tissues and organs. Useful for robotic extractions. Extraction protocol follows the manufacturer’s instructions.

4.2.3  Detection Procedures 4.2.3.1 One Step RT-PCR for the Detection of Type A Influenza Viruses This protocol is a modification of the method developed and described by Fouchier et al.82 for the detection of type A influenza viruses in samples of human and animal origin, including birds. According to previous field investigation on swabs of avian origin, the relative sensitivity was 95.6% (CI95 = 93.1–98.0) and the relative specificity was 96.3% (CI95 = 94.4–98.1) when compared to VI.32 In one other study on experimentally infected ducks48 the protocol provided 100% of relative sensitivity and 94% of

57

Avian Influenza Virus

relative specificity. In some cases, unspecific bands can be ­visualized on gel.52 Careful examination of the gel and use of proper controls and size-markers are therefore extremely important. Target: M gene Sample: RNA 5 µl in 25 µl of total reaction volume Reagents: One step RT-PCR (Applied Biosystems GeneAmp® Gold RNA PCR Core Kit Part No 4308207) Primers:82 Forward M52 C: 5′-CTT CTA ACC GAG GTC GAA ACG-3′ Reverse M253 R: 5′-AGG GCA TTT TGG ACA AAG/T CGT CTA-3′

a given thermocycler. The protocol has been tested mainly on H5 AI viruses belonging to the “Eurasian” lineage. Target: HA gene Sample: RNA 5 µl in 50 µl of total reaction volume Reagents: One step RT-PCR (Qiagen OneStep RT-PCR Kit cat # 210212) Primers:56 H5-kha-1: CCT CCA GAR TAT GCM TAY AAA ATT GTC H5-kha-3: TAC CAA CCG TCT ACC ATK CCY TG Note the inclusion of degenerate nucleotides indicated above in bold.

Reagent (Conc. Stock Solution)

Reagent

Final Concentration

Volume Required for one Reaction

/

4.7 µl

RNase-free water PCR buffer 5 × MgCl2 25 mM

1 × 2.5 mM

5 µl 2.5 µl

dNTPs mix 10 mM

1 mM

2.5 µl

DTT 100 mM

10 mM

2.5 µl

Primer M52 C 10 mM

0.3 mM

0.75 µl

Primer M253 R 10 mM

0.3 mM 10 U

0.75µl

Reverse transcriptase 50 U/µl

15 U

0.3 µl

Ampli Taq GOLD 5 U/µl Total volume Vortex the mix for few seconds. Aliquote 20 µl in 0.2-ml PCR tubes. RNA

2.5 U

0.5 µl

RNase inhibitor 20 U/µl

Final reaction volume

}

0.5 µl

20 µl 5 µl

RNase-free water PCR buffer 5 × from Qiagen OneStep RT-PCR Kit dNTPs mix 10 mM each (from Qiagen Kit)

Final Concentration

Volume Required for one Reaction (50 µl)

/

28.8 µl

1 × 

10 µl

0.4 mM each

2 µl

Primer H5-kha-1: 50 pmol/µl (50 µM)

1 µM

1 µl

Primer H5-kha-3: 50 pmol/µl (50 µM)

1 µM

1 µl

RNase inhibitor 40 U/µl (Promega) One Step RT-PCR Enzyme Mix (Qiagen Kit) Volume minus target

8U

0.2 µl 2 µl 45 µl

Volume extracted RNA

5 µl

Final reaction volume

50 µl

25 µl

Cycling conditions: One cycle of 42°C for 20 min; one cycle of 95°C for 5 min; 40 cycles of 94°C for 1 min, 55°C for 1 min, 72°C for 1 min; one cycle of 72°C for 10 min. Detection: Agarose gel 2 % or silver stained SDSPAGE 7% Expected amplified fragment: 244 bp.

Cycling conditions: One cycle of 50°C for 30 min; one cycle of 94°C for 15 min; 40 cycles of 94°C for 30 sec, 58°C for 1 min, and 68°C for 2 min; one cycle of 68°C for 7 min. Detection: Agarose gel 2 % or silver stained SDSPAGE 7% Expected amplified fragment: 300–320 bp.

4.2.3.2 One Step RT-PCR for the Detection of AI Viruses Belonging to the H5 Subtype This one step RT-PCR protocol has been partially validated in two consecutive ring trials performed in the EU and the results were published.56 According to the ring trials results, the protocol appears to be sensitive and useful for H5 AI virus detection in clinical specimens. Since the targeted region encompass the cleavage site segment of the HA gene segment, the subsequent sequence of the resulted amplified products will allow the determination of the pathotype. However, some results have revealed possible specificity problems. These include false positives with non-H5 AI specimens and/or multiple bands of similar size to the predicted amplicon. This may relate to the precise cycling conditions which are employed on

4.2.3.3 Two Step RT-PCR for the Subtype Specific Detection of H7 AI Virus This protocol consists in a two step RT-PCR first developed in The Netherlands and then selected for its good results among different tests during the AVIFLU European project.56 By this protocol, amplicon is detected conventionally by ­agarose gel electrophoresis with ethidium bromide staining or by SDS-PAGE with silver staining. The amplicon span the HA cleavage site so sequencing can provide pathotyping information, i.e., LPAI or HPAI. Target: HA gene Sample: RNA 5 µl in 50 µl of total reaction volume Primers:56 GK 7.3 5′-ATG TCC GAG ATA TGT TAA GCA-3′ GK 7.4 5′-TTT GTA ATC TGC AGC AGT TC-3′

58

Molecular Detection of Foodborne Pathogens

Step 1. Preparation of cDNA

Reagent

Final Concentration

RNA

/

Volume Required for one Reaction 10 µl

Primer GK 7.3 50 µM 2.5 µM 1 µl Heat to 95°C for 2 min and put the mix immediately on ice; then add the following reagents: RNase-free water / 3 µl

1 × 

M-MLV RT Buffer 5 ×  dNTPs mix 10 mM

0.5 mM each

RNase inhibitor 40 U/µl MMLV-RT 200 U/µl Final reaction volume

Probe:45 FAM M + 64 : FAM-5′-tca ggc ccc ctc aaa GCC GA-3′-TAMRA

Final Concentration

Volume Required for one Reaction

Probe FAM M + 64 1 µM

100 nM

2.5 µl

2 × QuantiTect Multiplex RT-PCR Master Mix

1 × 

12.5 µl

Reagent

4 µl

Primer M + 25F 5 µM

300 nM

1.5 µl

1 µl

Primer M-124R 5 µM QuantiTect Multiplex RT Mix

300 nM

1.5 µl

/

0.2 µl

/

1.8 µl

20 U

0.5 µl

100 U

0.5 µl 20 µl

Reverse transcriptase thermal parameters: 15 min at 37°C; 45 min at 42°C.

RNase-free water Total volume Vortex the mix for few seconds. Aliquote 20 µl per tube RNA Final reaction volume

Step 2. cDNA PCR amplification This protocol has been evaluated using the reagents contained in the AB Gene Kit cat#AB-0575/DC/LD/A:

Final Concentration

Volume Required for one Reaction

/

18 µl

1 × 

25 µl

Primer GK 7.3 50 µM

1 µM

1 µl

Primer GK 7.4 50 µM Total volume

1 µM

1 µl

Reagent RNase-free water 2 × Reddy Mix PCR Master Mix with dNTPs

cDNA Final reaction volume

}

45 µl 5 µl 50 µl

Cycling conditions: one cycle of 94°C for 30 min; 35 cycles of 94°C for 30 sec, 52°C for 30 sec, 72°C for 45 sec; one cycle of 72°C for 4.15 min. Detection: agarose gel 2% or silver stained SDSPAGE 7% Expected amplified fragment: 200–220 bp. 4.2.3.4 Detection of Type A Influenza Virus by Qualitative Real Time PCR (M gene) The protocol uses the probe-primer set previously evaluated by Spackman et al.45 The basic procedure has been previously evaluated by different authors.45,56 In the one step rRT-PCR protocol described below the QuantiTect Multiplex RT-PCR Kit (Qiagen Cod. 204643) has been adopted. Reagents: QuantiTect Multiplex RT-PCR Kit (Qiagen Cod. 204643) Primers:45 Forward M + 25: AGA TGA GTC TTC TAA CCG AGG TCG Reverse M-124: TGC AAA AAC ATC TTC AAG TCT CTG

}

20 µl 5 µl 25 µl

Cycling conditions: This protocol was developed on AB7300 (Applied Biosystems) and Rotorgene 6000 (Corbett) real time platforms. Other laboratories using different instrumentation platforms should first critically and carefully examine these cycling conditions as they may not perform optimally on other instruments. Hence cycling temperatures, times and ramp speeds may all need to be modified. Thermal parameters: one cycle of 50°C for 20 min; one cycle of 95°C for15 min; 40 cycles of 94°C for 45 sec, 60°C for 45 sec. 4.2.3.5 Detection of Type A Influenza Viruses of H5 HA Subtype by Qualitative One Step Real Time RT-PCR Similarly to the H7 real time PCR protocol (Section 4.2.3.6), this protocol has been developed and properly validated recently.46 The procedure can be coupled with H7 detection within the same real time PCR run, providing a fast and sensitive method for the detection of notifiable AI subtypes in poultry. The protocol has been validated on a wide variety of H5/H7 isolates of the Eurasian lineage, including the recent H5/H7 LPAI viruses circulating in poultry and wild birds as well as the H5N1 HPAI viruses circulating in Eastern and Central Asia, Middle East, Europe and Africa. The protocol was developed using the QuantiTect Multiplex RT-PCR Kit (Qiagen Cod. 204643). Reagents: QuantiTect Multiplex RT-PCR Kit (Qiagen Cod. 204643). Primers:46 Forward H5 F : TTA TTC AAC AGT GGC GAG Reverse H5 R: CCA KAA AGA TAG ACC AGC Note degenerate nucleotides indicated in bold. Probe:46 FAM H5: FAM-5′-ccc tag cac tgg caa tca tg-3′-TAMRA

59

Avian Influenza Virus

Reagent (Conc. Stock Solution)

Final Concentration

Probe FAM H5 1 µM

150 nM

2 × QuantiTect Multiplex RT-PCR Master Mix

1 × 

Volume Required for one Reaction 3.75 µl 12.5 µl

Primer H5F 5 µM

300 nM

1.5 µl

Primer H5Rnew 5 µM QuantiTect Multiplex RT Mix

300 nM

1.5 µl

RNase-free water Total volume Vortex the mix for few seconds. Aliquote 20 µl per tube RNA

0.2 µl /

}

0.55 µl 20 µl 5 µl 25 µl

Final reaction volume

Cycling conditions: This protocol was evaluated on AB7300 (Applied Biosystems) and Rotorgene 6000 (Corbett) real time platforms. Other laboratories using different instrumentation platforms should first critically and carefully examine these cycling conditions as they may not perform optimally on other instruments. Hence cycling temperatures, times and ramp speeds may all need to be modified. Thermal parameters: one cycle of 50°C for 20 min; one cycle of 95°C for 15 min; 40 cycles of 94°C for 45 sec, 54°C for 45 sec. 4.2.3.6 Detection of Type A Influenza Viruses of H7 HA Subtype by Qualitative Real Time PCR 46 Primers: Forward H7 F: TTT GGT TTA GCT TCG GG Reverse H7 R: GAA GAM AAG GCY CAT TG Note degenerate nucleotides indicated in bold. Probe:46 VIC H7: VIC-5′-CAT CAT GTT TCA TAC TTC TGG CCA T-3 ′-TAMRA

Reagent Probe VIC H7 1 µM 2 × QuantiTect Multiplex RT-PCR Master Mix

Final Concentration

Volume Required for one Reaction

150 nM

3.75 µl

1 × 

12.5 µl

Primer H7F 10 µM

300 nM

0.75 µl

Primer H7R 10 µM QuantiTect Multiplex RT Mix

900 nM

2.25 µl

/

0.55 µl

RNase-free water Total volume Vortex the mix for few seconds. Aliquote 20 µl per tube RNA Final reaction volume

real time platforms. Other laboratories using different instrumentation platforms should first critically and carefully examine these cycling conditions as they may not perform optimally on other instruments. Hence cycling temperatures, times and ramp speeds may all need to be modified. Thermal parameters: one cycle of 50°C for 20 min; one cycle of 95°C for 15 min; 40 cycles of 94°C for 45 sec, 54°C for 45 sec.

0.2 µl

}

20 µl 5 µl 25 µl

Cycling conditions: This protocol was evaluated on AB7300 (Applied Biosystems) and Rotorgene 6000 (Corbett)

4.3 Conclusions and Future Perspectives The increasing importance of AI viruses for the veterinary and medical sciences in the last decade has contributed to a tremendous increase in actions related to this infection with the aim to better understand the pathogenicity and virulence mechanisms of these viruses and to develop better diagnostic tools for their detection. Taking advantages of the new technologies available nowadays for the diagnosis of infectious diseases, an impressive number of scientific papers has been published describing the application of technologies, such as NASBA, LAMP-PCR, or microarrays, to the detection of the viruses. Further, well established technologies, such as RT-PCR or real time PCR have been improved and applied for this infection. As for many other infectious and contagious diseases, it has been realized that rapidity in the detection and characterization of the responsible microorganism is essential for the disease control. The key issues related to the major changes in the field of AI diagnosis in the last 10–15 years, can be summarized into two main points: improved rapidity and flexibility. If compared to the classical VI and typing methods, molecular technologies have probably not improved significantly the diagnostic sensitivity or specificity. Rather, they have allowed the detection of the causative agent, in association with its typing, subtyping, and with the characterization of its molecular determinant of pathogenicity in a more time-effective and flexible manner. Importantly, the molecular tests have made the screening of large susceptible populations, such as poultry or wild birds, sustainable and cost-effective for many countries. Many molecular protocols are currently available for AI testing, but only few of these have been fully and properly validated at present. Provision and harmonization of molecular test validation procedures is therefore necessary to expand the number of validated protocols, getting to more standardized laboratory results. With very few exceptions, the vast majority of the available protocols have been developed and tested for the detection of AI viruses in clinical specimens, from alive or dead animals. To better perform epidemiological investigations, as well as to better address public health and food safety issues, it would be desirable to expand the application of the molecular tests to environmental specimens, such as water, soil, surfaces, etc., and to poultry-derived products such as meat, feathers, and eggs. Thus, the development of new protocols or the validation of the existing testing procedures on these materials appears to be necessary.

60

References











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Molecular Detection of Foodborne Pathogens 23. Guo, Y., Li, J. and Cheng, X. Discovery of men infected by avian influenza A (H9N2) virus. Zhonghua Shi Yan He Lin Chuang Bing Du Xue Za Zhi, 13, 105, 1999. 24. Puzelli, S. et al. Serological analysis of serum samples from humans exposed to avian H7 influenza viruses in Italy between 1999 and 2003. J. Infect. Dis., 192, 1318, 2005. 25. Hosseini, M. et al. Seroprevalence of H9N2 antibody in poultry farm and slaughter house workers of Iran using HI test. Proceedings of the 13th World Association of Veterinary Diagnosticians Symposium, 11–14 November 2007, Melbourne, Australia. 26. Subbarao, K. et al. Characterization of an avian influenza A (H5N1) virus isolated from a child with a fatal respiratory ­illness. Science, 279, 393, 1998. 27. Peiris, J.S et al. Re-emergence of fatal human influenza A subtype H5N1 disease. Lancet, 21, 363, 617, 2004. 28. Koopmans, M. et al. Transmission of H7N7 avian influenza A virus to human beings during a large outbreak in commercial poultry farms in the Netherlands. Lancet, 363, 587, 2004. 29. Tweed, S.A. et al. Human illness from avian influenza H7N3, British Columbia. Emerg. Infect. Dis., 10, 2196, 2004. 30. OIE (World Organization for Animal Health), Highly pathogenic avian influenza. In: Manual of Diagnostic Tests and Vaccines for Terrestrial Animals, 5th Edition. Office International des Epizooties, Paris, France, 258, 2004. 31. EC (European Commission), Commission Decision 2006/437/ EC of 4 August 2006 approving a diagnostic manual for avian influenza as provided for in Council Directive 2005/94/EC [notified under document number C (2006) 3477], 2006. Available at: http://eur-lex.europa.eu/LexUriServ/site/en/oj/2006/l_237/ l_23720060831en00010027.pdf 32. Cattoli, G. et al. Comparison of three rapid detection systems for type A influenza virus on tracheal swabs of experimentally and naturally infected birds. Avian Pathol., 33, 432, 2004. 33. Woolcock, P.R. and Cardona, C.J. Commercial immunoassay kits for the detection of influenza virus type A: evaluation of their use with poultry. Avian Dis., 49, 477, 2005. 34. Cattoli, G. and Capua, I. Diagnosing avian influenza in the framework of wildlife surveillance efforts and environmental samples. J. Wildlife Dis., 43, S35, 2007. 35. Higgins, D.A. Precipitating antibodies of the duck (Anas plathyrhyncos). Comp. Biochem. Physiol., 93B, 135, 1989. 36. Cattoli, G. and Capua, I. Molecular diagnosis of avian influenza during an outbreak. Dev. Biol. (Basel), 124, 99, 2006. 37. Collins, R.A. et al. A NASBA method to detect high- and lowpathogenicity H5 avian influenza viruses. Avian Dis., 47, 1069, 2003. 38. Lau, L.T. et al. Nucleic acid sequence-based amplification methods to detect avian influenza virus. Biochem. Biophys. Res. Commun., 313, 336, 2004. 39. Imai, M. et al. Development of H5-RT-LAMP (loop-mediated isothermal amplification) system for rapid diagnosis of H5 avian influenza virus infection. Vaccine, 24, 6679, 2006. 40. Jayawardena, S. et al. Loop-mediated isothermal amplification for influenza A (H5N1) virus. Emerg. Infect. Dis., 13, 899, 2007. 41. Pourmand, N. et al. Rapid and highly informative diagnostic assay for H5N1 influenza viruses. PLoS One, 20,1e95, 2006. 42. Ellis, J.S. et al. Design and validation of an H5 TaqMan real-time one-step reverse transcription-PCR and confirmatory assays for diagnosis and verification of influenza A virus H5 infections in humans. J. Clin. Microbiol., 45, 1535, 2007.

Avian Influenza Virus 43. Forster, J.L. et al. The effect of sample type, temperature and RNAlater trade mark on the stability of avian influenza virus RNA. J. Virol. Methods, 149, 190, 2008. 44. Dybkaer, K., et al. Application and evaluation of RT-PCRELISA for the nucleoprotein and RT-PCR for detection of low-pathogenic H5 and H7 subtypes of avian influenza virus. J. Vet. Diagn. Invest., 16, 51, 2004. 45. Spackman, E. et al. 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, 2002. 46. Monne, I. et al. Development and validation of a one step real time PCR assay for the simultaneous detection of H5, H7 and H9 subtype avian influenza viruses. J. Clin. Microbiol., 46, 1769, 2008. 47. Lee, C.W. and Suarez, D.L. Application of real-time RT-PCR for the quantitation and competitive replication study of H5 and H7 subtype avian influenza virus. J. Virol. Methods, 119, 151, 2004. 48. Foni, E. et al. Detection of influenza A virus by RT-PCR and standard methods in experimental infection of Ducks. New Microbiol., 28, 31, 2005. 49. Xing, Z. et al. Realtime RT-PCR assay unable to detect H7 subtype avian influenza viruses isolated from wild birds. J. Clin. Microbiol., 46,1844, 2008. 50. Bogner, P. et al. A global initiative on sharing avian flu data. Nature, 442, 981, 2006. 51. OIE (World Organization for Animal Health). Quality Standard and Guidelines for Veterinary Laboratories: Infectious Diseases, 2nd Edition. Office International des Epizooties, Paris, France, 2008, Available online at http://www.oie.int. 52. Martì, N.B. et al. False positive results obtained by following a commonly used reverse transcription-PCR protocol for detection of Influenza A virus. J. Clin. Microbiol., 44, 3845, 2006. 53. Das, A. et al. Development of an internal positive control for rapid diagnosis of avian influenza virus infections by real time reverse transcription-PCR with lyophilized reagents. J. Clin. Microbiol., 44, 3065, 2006. 54. Slomka, M.J. et al. Validated real time reverse transcriptasepolymerase chain reaction and its application in H5N1 outbreaks in 2005–2006. Avian Dis., 50, 373, 2007. 55. Suarez, D.L. Evolution of avian influenza viruses. Vet. Microbiol., 74, 15, 2000. 56. Slomka, M.J. et al. Identification of sensitive and specific avian influenza polymerase chain reaction methods through blind ring trials organized in the European Union. Avian Dis., 51, 227, 2007. 57. Hoffmann, B. et al. Rapid and highly sensitive pathotyping of avian influenza A H5N1 virus by using real-time reverse transcription-PCR. J. Clin. Microbiol., 45, 600, 2007. 58. Banks, J. et al. Phylogenetic analysis of influenza A viruses of H9 haemagglutinin subtype. Avian Pathol., 29, 353, 2000. 59. Lee, M-S. et al. Identification and subtyping of avian influenza viruses by reverse transcription-PCR. J. Virol. Methods, 97, 13, 2001. 60. Phipps, L.P., Essen, S.C. and Brown, I.H. Genetic subtyping of influenza A viruses using RT-PCR with a single set of primers based on conserved sequences within the HA2 coding region. J. Virol. Methods, 122, 119, 2004. 61. Wang, R. et al. Examining the hemagglutinin subtype diversity among wild duck-origin influenza A viruses using ethanol-fixed cloacal swabs and a novel RT-PCR method. Virology, 375, 182, 2008.

61 62. Guan, J. et al. Development of methods for detection and quantification of avian influenza and newcastle disease viruses in compost by real-time reverse transcription polymerase chain reaction and virus isolation. Poultry Sci., 87, 838, 2008. 63. Capua, I. et al. Increased resistance of vaccinated turkeys to experimental infection with an H7N3 low-pathogenicity avian influenza virus. Avian Pathol., 33, 158, 2004. 64. Keawcharoen, J. et al. Wild ducks as long-distance vectors of highly pathogenic avian influenza virus (H5N1). Emerg. Infect. Dis., 14, 600, 2008. 65. Webster, R.G., et al. Intestinal influenza: replication and characterization of influenza viruses in ducks. Virology, 84, 268, 1978. 66. Tian, G. et al. Protective efficacy in chickens, geese and ducks of an H5N1-inactivated vaccine developed by reverse genetics. Virology, 341, 153, 2005. 67. Rimmelzwaan, G.F., Osterhaus, A.D. and Kuiken, T. Influenza A virus (H5N1) infection in cats causes systemic disease with potential novel routes of virus spread within and between hosts. Am. J. Pathol., 168, 176, 2006. 68. Mase, M. et al. Experimental assessment of the pathogenicity of H5N1 influenza A viruses isolated in Japan. Avian Dis., 49, 582, 2005. 69. Swayne, D.E. and Beck, J.R. Experimental study to determine if low-pathogenicity and high-pathogenicity avian influenza viruses can be present in chicken breast and thigh meat following intranasal virus inoculation. Avian Dis., 49, 81, 2005. 70. Muramoto, Y. et al. Highly pathogenic H5N1 influenza virus causes coagulopathy in chickens. Microbiol. Immunol., 50, 73, 2006. 71. Kishida, N. et al. Pathogenicity of H5 influenza viruses for ducks. Arch. Virol., 150, 1383, 2005. 72. Kishida, N. et al. Co-infection of Staphylococcus aureus or Haemophilus paragallinarum exacerbates H9N2 influenza A virus infection in chickens. Arch. Virol., 149, 2095, 2004. 73. Toffan, A. et al. Conventional inactivated bivalent H5/H7 vaccine prevents viral localisation in muscles of turkeys infected experimentally with LPAI and HPAI H7N1 isolates. Avian Pathol., 37, 407, 2008. 74. Tumpey, T.M. et al. Characterization of highly pathogenic H5N1 avian influenza A virus isolated from duck meat. J. Virol., 76, 6344, 2002. 75. Beato, M.S. et al. A conventional, inactivated oil emulsion vaccine suppresses shedding and prevents viral meat colonisation in commercial (Pekin) ducks challenged with HPAI H5N1. Vaccine, 25, 4064, 2007. 76. Beato, M.S. et al. Isolation and characterization of an H10N7 avian influenza virus from poultry carcasses smuggled from China into Italy. Avian Pathol., 35, 400, 2006. 77. Beato, M.S. et al. Inactivation of avian influenza viruses by nucleic acid extraction reagents. Proceedings of the 13th World Association of Veterinary Diagnosticians Symposium, 11–14 November 2007, Melbourne, Australia. 78. Evers, D.L., Slemons, R.D. and Taubenberger, J.K. Effect of preservative on recoverable RT-PCR amplicon length from influenza A virus in bird feces. Avian Dis., 51, 965, 2007. 79. Taylor, H.R. and Turner A.J. A case report of fowl plague keraconjunctivitis. Brit. J. Ophthalmol., 61, 86, 1977. 80. WHO Laboratory Biosafety Manual, 3rd Edition, 2004. Available online at http://www.who.int/csr/resources/ publications/biosafety/WHO_CDS_CSR_LYO_2004_11/en/. 81. WHO Laboratory Biosafety Guidelines for Handling Specimens Suspected of Containing Avian Influenza A Virus, 2005. Available online at http://www.who.int/csr/disease/ avian_influenza/guidelines/handlingspecimens/en/.

62 82. Fouchier, R.A.M. et al. Detection of influenza A viruses from different species by PCR amplification of conserved sequences in the matrix gene. J. Clin. Microbiol., 38, 4096, 2000. 83. Starick, E., Römer-Oberdörfer, A. and Werner, O. Type- and subtype-specific RT-PCR assays for avian influenza A viruses (AIV). J. Vet. Med. B. Infect. Dis. Vet. Public Health, 47, 295, 2000. 84. Farkas, T. et al. Rapid and simultaneous detection of avian influenza and newcastle disease viruses by duplex polymerase chain reaction assay. Zoonoses Public Health., 54, 38, 2007. 85. Malik, Y.S., Patnayak, D.P. and Goyal, S.M. Detection of three avian respiratory viruses by single-tube multiplex reverse transcription-polymerase chain reaction assay. J. Vet. Diagn. Invest., 16, 244, 2004. 86. Chaharaein, B. et al. Detection of H5, H7 and H9 subtypes of avian influenza viruses by multiplex reverse transcriptionpolymerase chain reaction. Microbiol. Res., doi:10.1016/j. micres.2007.01.001. 87. Xie, Z. et al. A multiplex RT-PCR for detection of type A influenza virus and differentiation of avian H5, H7, and H9 hemagglutinin subtypes. Mol. Cell. Probes, 20, 245, 2006. 88. Thontiravong, A. et al. The single-step multiplex reverse transcription- polymerase chain reaction assay for detecting H5 and H7 avian influenza A viruses, Tohoku. J. Exp. Med., 211, 75, 2007. 89. Ng, L.F. et al. Specific detection of H5N1 avian influenza A virus in field specimens by a one-step RT-PCR assay. BMC Infect. Dis., 2, 6, 2006. 90. Di Trani, L. et al. A sensitive one-step real-time PCR for detection of avian influenza viruses using a MGB probe and an internal positive control. BMC Infect. Dis., 6, 87, 2006.

Molecular Detection of Foodborne Pathogens 91. Kiss, I. et al. Application of real-time RT-PCR utilising lux (light upon extension) fluorogenic primer for the rapid detection of avian influenza viruses. Acta Vet. Hung., 54, 525, 2006. 92. Karlsson, M. et al. A real-time PCR assay for the monitoring of influenza A virus in wild birds. J. Virol. Methods, 144, 27, 2007. 93. Payungporn, S. et al. Single step multiplex real-time RT-PCR for H5N1 influenza A virus detection. J. Virol. Methods, 131, 143, 2006. 94. Li, P.Q. et al. Development of a multiplex real-time polymerase chain reaction for the detection of influenza virus type A including H5 and H9 subtypes. Diagn. Microbiol. Infect. Dis., 61, 192, 2008. 95. Ong, W.T. et al. Development of a multiplex real-time PCR assay using SYBR Green 1 chemistry for simultaneous detection and subtyping of H9N2 influenza virus type A. J. Virol. Methods, 144, 57, 2007. 96. Rossi, J., Cramer, S. and Laue, T. Sensitive and specific detection of influenza virus A subtype H5 with real-time PCR. Avian Dis., 51, 387, 2007. 97. Wu, C. et al. A multiplex real-time RT-PCR for detection and identification of influenza virus types A and B and subtypes H5 and N1. J. Virol. Methods, 148, 81, 2008. 98. Lu, Y.Y et al. Rapid detection of H5 avian influenza virus by TaqMan-MGB real-time RT-PCR. Lett. Appl. Microbiol., 46, 20, 2008. 99. Ng, K.O.E., et al. Influenza A H5N1 detection. Emerg. Infect. Dis., 11, 1303, 2005. 100. Agüero, M. et al. A real-time TaqMan RT-PCR method for neuraminidase type 1 (N1) gene detection of H5N1 Eurasian strains of avian influenza virus. Avian Dis., 51 (Suppl 1), 378, 2007.

5 Hepatitis A and E Viruses Hiroshi Ushijima, Pattara Khamrin Aino University

Niwat Maneekarn

Chiang Mai University

Contents 5.1 Introduction ........................................................................................................................................................................ 63 5.1.1 Hepatitis A Virus (HAV)......................................................................................................................................... 63 5.1.1.1 Overview of Hepatitis A Virus Infection and Pathogenesis..................................................................... 63 5.1.1.2 Hepatitis A Virion, Genome Organization, and Proteins......................................................................... 63 5.1.1.3 HAV Classification and Genetic Diversity................................................................................................ 64 5.1.1.4 Foodborne and Waterborne HAV.............................................................................................................. 64 5.1.1.5 Diagnosis of HAV..................................................................................................................................... 64 5.1.2 Hepatitis E Virus (HEV)......................................................................................................................................... 65 5.1.2.1 Overview of Hepatitis E Virus Infection and Pathogenesis...................................................................... 65 5.1.2.2 Hepatitis E Virion, Genome Organization, and Proteins.......................................................................... 66 5.1.2.3 HEV Classification and Genetic Diversity................................................................................................ 66 5.1.2.4 Foodborne and Waterborne HEV............................................................................................................. 66 5.1.2.5 Diagnosis of HEV..................................................................................................................................... 66 5.2 Methods............................................................................................................................................................................... 67 5.2.1 Reagents and Equipments........................................................................................................................................ 67 5.2.2 Sample Collection and Preparation ........................................................................................................................ 68 5.2.2.1 HAV Specimen Collection and RNA Extraction ..................................................................................... 68 5.2.2.2 HEV Specimen Collection and RNA Extraction ..................................................................................... 68 5.2.3 Detection Procedures . ............................................................................................................................................ 68 5.2.3.1 Detection of HAV . ................................................................................................................................... 68 5.2.3.2 Detection of HEV . ................................................................................................................................... 70 5.3 Conclusions and Future Perspectives.................................................................................................................................. 71 References.................................................................................................................................................................................... 71

5.1  Introduction 5.1.1 Hepatitis A Virus (HAV) 5.1.1.1 Overview of Hepatitis A Virus Infection and Pathogenesis Viral hepatitis is a major health concern worldwide with higher incidence in developing countries than in the developed countries.1,2 Hepatitis A virus (HAV) is one of the etiologic agents of acute viral hepatitis. Most infection occur in children and generally are self-limiting.1 Transmission of HAV is primarily via the fecal-oral route, either by contact with an infected person or by ingestion of contaminated food and water.1 In the industrialized countries, due to improvements of public health and socioeconomic conditions, there has been a shift of HAV infection toward a higher age with an increase of hospitalized and severe cases associated with outbreaks.3 The illness severity is age-dependent. Generally,

HAV infection in children is asymptomatic and rarely develops jaundice, whereas in older children and adults are symptomatic infection with a wide range of clinical manifestations from mild and anicteric infection to severe and fulminant hepatic failure (FHF).1 Infected people can excrete HAV in feces for about 3 months or longer, and viremia is detectable by reverse transcription-polymerase chain reaction (RT-PCR) in the majority of patients at the onset of symptoms and can persist for several weeks after aminotransferase peak.4 Viral replication occurs primarily within hepatocytes, and then the viruses are secreted into bile through the bile ducts which results in large amount of virus being shed in the feces.5 5.1.1.2 Hepatitis A Virion, Genome Organization, and Proteins HAV is classified as the only member of the genus Hepatovirus within the Picornaviridae family.6,7 It is a 63

64

spherical, icosahedral symmetry, nonenveloped RNA virus. The virus particle is 27–32 nm in diameter with a positive single-stranded RNA genome of 7.5 kb.1 The viral genome composed of 5′ nontranslated region (NTR), structural protein regions, nonstructural protein regions, 3′ NTR, and followed by a short poly(A) tail.8 A single large open reading frame (ORF) of HAV genome can be divided into three distinct functional protein-encoded regions termed P1, P2, and P3. The P1 region encodes the capsid polypeptides VP1–VP4. The P2 and P3 regions encode the nonstructural polypeptides which are necessary for virus replication.8 5.1.1.3 HAV Classification and Genetic Diversity The HAV strains isolated from various parts of the world constitute a single serotype. However, genetic variability between strains allows the classification of HAV into six different genotypes based on phylogenetic analysis of nucleotide sequences in the VP1/P2A region.9,10 Based on this region, the HAV strains that differ from each other at least 15% of nucleotide sequences are considered to be different genotypes, while the strains that their nucleotide sequence differ over 7–7.5% belong to different HAV subgenotypes.9 Of six HAV genotypes, genotype I, II, and III are associated with human infections, and are further divided into subgenotype IA, IB, IIA, IIB, IIIA, and IIIB, respectively. The remaining three genotypes IV, V, and VI are represented by strains of simian HAV.9,10 From the epidemiological data, HAV genotypes I and III are the vast majority of human strains which comprise more than 80% of strains circulating worldwide.1,9,11 5.1.1.4  Foodborne and Waterborne HAV Foodborne and waterborne viral infections are increasingly recognized as the causes of illness in humans. HAV is one of the leading causes of foodborne and waterborne viral infections. It can be transmitted from person-to-person, or indirectly via food and water contaminated with virus-containing feces. HAV has previously been isolated directly from food or environmental sources. Salad vegetables, soft fruits, green onion, strawberry, lettuce, clam, shellfish, and oyster samples have been reported to be the sources of HAV infections.12–17 A large epidemic outbreak of HAV, reported in 1988, was attributed to the ingestion of raw clams and caused illness in 300,000 persons in Shanghai, China.18 Moreover, waterborne outbreaks of HAV associated with HAV contamination in water supply have also been reported from several countries. HAV has also been detected in river, canal, ground/tap water and sewage.12,16,19–21 5.1.1.5 Diagnosis of HAV For the detection of HAV genomic RNA, several genome segments have been amplified by different sets of primer pairs. The genomic regions which have been used widely to detect

Molecular Detection of Foodborne Pathogens

and to define HAV genotypes, included the junction of VP3/ VP1 region.17,22–25 the junction of VP1/P2A region,4,24–28 the entire VP1 region,29 and the junction of 3C/3D region.30 The primers used for the detection of HAV are summarized in Table 5.1. Currently, the genomic region that most commonly used for the detection and identification of HAV genotypes is the VP1/P2A junction region. To facilitate the molecular analysis of HAV, the amplification of their genomic RNA and sequencing of DNA amplicon should be performed and HAV genotypes are identified based on sequence analysis. Sequence variation within the VP1/P2A junction has defined six HAV genotypes and two subtypes within genotypes I, II, and III.9–11 In contrast to most picornaviruses, HAV of human origin replicates poorly in cell cultures with a relatively low concentration of viruses and viral antigen being produced into the cultured supernatant. For these reasons, development of a number of RT-PCR-based assays that enable the rapid and specific detection of small amount of viral nucleic acid in environmental sources, food samples, and clinical specimens have been developed recently. Several kinds of clinical samples have been used for the detection of HAV genome, including stool, serum, saliva, and liver suspension.9,26,31 In addition, HAV has also been detected by other sensitive molecular techniques such as restriction fragment length polymorphism (RFLP),32 Southern blotting,33 real-time RT-PCR,34 and reverse transcription loop-mediated isothermal amplification assay (RT-LAMP).35 Currently, the amplification of viral RNA by RT-PCR is the most sensitive and widely used method for the detection of foodborne HAV RNA.15,16,21,25,36 The efficiency of the extraction methods for HAV RNA from clinical specimens is of great important for molecular diagnosis. Therefore, choosing appropriate RNA extraction methods is a critical step for a successful and valid use of PCR amplification of viral genome in clinical samples. Traditionally, proteinase K digestion and guanidinium isothiocyanate (GTC)-phenol-chloroform extraction method followed by ethanol precipitation has been widely used to extract RNA from serum or stool samples.37 Recently, numerous protocols for RNA extraction, i.e., GTC-silica method, antigen-capture method, and magnetic beads coated with anti-HAV have been used for the isolation of HAV RNA and to separate viral genome from the potential inhibitors of RT-PCR reaction that might exist in the clinical samples.9,22,31,37–41 The total genomic RNA of HAV can also be easily isolated from clinical samples with high sensitivity and reproducibility by using commercially available RNA extraction kits, i.e., Trizol LS® Reagent (Invitrogen Life Technology, Carlsbad, CA)30,41 or QIAamp Viral RNA Mini Kit (Qiagen, Hilden, Germany).27,31,35,42 Recently, the use of the QIAamp Viral RNA Mini Kit (Qiagen) has been reported to be an efficient method for extraction of HAV genomic RNA with a detection limit of 6 × 103 copies/ml in clinical samples.31

65

Hepatitis A and E Viruses

Table 5.1 Oligonucleotide Primers for the Detection and Sequencing of HAV from Clinical Specimens Primer Name

Region

Sequence 5′–3′

Position

Sense

VP1-4

VP3/VP1

First PCR

CGT TGC TTC CCA TGT CAG AG

2115–2135

+

VP1-5

VP3/VP1

First PCR

GAC CTT CCC ATA AAC TTG TAG

2483–2462

–

VP1-2

VP3/VP1

Second PCR

GTT TTG CTC CTC TTT ATC ATG CTA TG

2168–2194

+

VP1-1

VP3/VP1

Second PCR

GGA AAT GTC TCA GGT ACT TTC TTT G

2415–2390

–

BR-5

VP1/P2A

First PCR

TTG TCT GTC ACA GAA CAA TCA G

2950–2972

+

BR-9

VP1/P2A

First PCR

AGT CAC ACC TCT CCA GGA AAA CTT

3310–3286

–

RJ-3

VP1/P2A

Second PCR

TCC CAG AGC TCC ATT GAA

2984–3002

+

BR-6

VP1/P2A

Second PCR

AGG AGG TGG AAG CAC TTC ATT TGA

3217–3193

–

–

VP3/VP1

First PCR

GTG AAT GTT TAT CTT TCA GCA AT

2133–2155

+

–

VP3/VP1

First PCR

GAT CTG ATG TAT GTC TGG ATT CT

2429–2451

–

–

VP3/VP1

Second PCR

GCT CCT CTT TAT CAT GCT ATG GAT

2172–2195

+

–

VP3/VP1

Second PCR

CAG GAA ATG TCT CAG GTA CTT TCT

2392–2415

–

–

VP1/P2A

First PCR

GAC AGA TTC TAC ATT TGG ATT GGT

2870–2893

+

–

VP1/P2A

First PCR

CCA TTT CAA GAG TCC ACA CAC T

3360–3381

–

–

VP1/P2A

Second PCR

CTA TTC AGA TTG CAA ATT ACA AT

2896–2981

+

–

VP1/P2A

Second PCR

AAC TTC ATT ATT TCA TGC TCC T

3268–3289

–

+2897

VP1/P2A

First PCR

CTA TTC AGA TTG CAA ATT AYA AT

2897–2819

+

–3288

VP1/P2A

First PCR

AAY TTC ATY ATT TCA TGC TCC T

3267–3288

–

+2949

VP1/P2A

Second PCR

TAT TTG TCT GTY ACA GAA CAA TCA G

2949–2973

+

–3192

VP1/P2A

Second PCR

AGG RGG TGG AAG YAC TTC ATT TGA

3169–3192

–

A

VP3/VP1

First PCR

GTT TTG CTC CTC TTT ATC ATG CTA TG

2167–2192

+

B

VP3/VP1

First PCR

GGA AAT GTC TCA GGT ACT TTC TTT G

2389–2414

–

A

VP3/VP1

Second PCR

GTT TTG CTC CTC TTT ATC ATG CTA TG

2167–2192

+

C

VP3/VP1

Second PCR

TCC TCA ATT GTT GTG ATA GC

2358–2377

–

HA027

3C/3D

First PCR

TTT GGW GTT GGD GAG AAR AAT GG

5318–5340

+

HA030

3C/3D

First PCR

AAG CGT TTT GGA GAC YAC ATT C

5969–5989

–

HA028

3C/3D

Second PCR

GTG TKA GRT GGG TYA TGA ATG C

5343–5364

+

HA029

3C/3D

Second PCR

GAC YAC ATT CAT TGA ACA YTG AG

5955–5977

–

5.1.2 Hepatitis E Virus (HEV) 5.1.2.1 Overview of Hepatitis E Virus Infection and Pathogenesis HEV is widely spread in many tropical and subtropical countries where sanitary infrastructure remains inadequate.43 Clinical manifestations of HEV infection are similar to those of HAV infection and encompass a wide spectrum of symptoms. Typical HEV symptoms include jaundice, hepatomegaly, dark- or tea-colored urine, anorexia, diarrhea, vomiting, and fever.44,45 The illness is usually self-limiting and typically lasts for about 1–4 weeks and infection does not progress to chronicity. The disease is slightly more severe than HAV infection, but still has a low mortality in the general population with the rate of about 1–4%.44,46 However, a few patients develop a prolonged clinical illness

PCR Product

Reference

369 bp

24

247 bp

24

360 bp

24, 26, 27

234 bp

24, 26, 27

319 bp

25

244 bp

25

512 bp

25

394 bp

25

392 bp

4, 28

244 bp

9, 28

248 bp

23

211 bp

23

672 bp

30

635 bp

30

24 24

24, 26, 27 24, 26, 27

25 25

25 25

4, 28 9, 28

23 23

30 30

and severe complications with a high mortality rate associated with infection of pregnant women (15–20%), particularly in the third trimester of pregnancy. Causes of death among pregnant women with HEV infection include hepatic encephalopathy and disseminated intravascular coagulation.43,47–50 Incubation period of HEV infection ranges from 15 to 60 days.51–53 HEV antigen first appears in the liver and this is typically followed by viremia.54 Virus accumulates at a relatively high concentration in bile and is subsequently shed in the feces. Viremia and fecal shedding are detected prior to liver abnormalities. Liver function tests are indicative of hepatic inflammation. Laboratory tests of enzyme level abnormalities including, high peak of alanine aminotransferase (ALT), aspartate aminotransferase (AST), gammaglutamyl aminotransferase (GGT), and serum alkaline phosphatase (SAP). Experimental studies

66

of HEV infections in human volunteers demonstrated that liver enzyme elevations occurred at 4–5 weeks after oral ingestion and persisted for 3–13 weeks.51,53 Virus excretions in feces occurred approximately at 2–4 weeks. In addition, HEV RNA could be detected in serum around the first 2–3 weeks after exposure. Antibodies to HEV were first detected at about 6 weeks after infection and remained detectable up to 2 years.51,53 The HEV pathogenesis is not well understood. Several studies demonstrated that HEV is transmitted through fecal-oral contact or consumption of contaminated food or water.43,55–57 The site of the primary replication has not been identified, but it is presumed to be in the intestinal tract. The route and mechanism by which the virus reaches the liver from the intestinal tract after consumption of the virus remain unclear. However, possible mechanism of HEV associated disease may be due to (i) liver cell injury caused by either an immunologic mechanism and/or direct cytopathic effects of the virus on the liver cells, and (ii) viral antigenantibody complex mediated vasculitis/glomerulonephritis.47 HEV particle is almost certainly resistant to the change of pH between alkaline and acidic. Therefore, it can survive in the gastrointestinal tract and cause infection, but does not seem to tolerate exposure to high salt concentration, including cesium chloride. Since the main mode of transmission of HEV is the fecal-oral route, prevention of HEV infection, therefore, relies primarily on the provision of clean drinking water, improving personal hygiene, and avoiding consumption of raw, uncooked meat, and vegetables. Study of thermal stability of HEV suggests that it is less stable than HAV.58 5.1.2.2 Hepatitis E Virion, Genome Organization, and Proteins Initially, HEV had been classified as a member of the Picornaviridae family. Later, it was shown that HEV was antigenically and biophysically unrelated to the picornaviruses. For these reasons, HEV has been reclassified as the only member of the genus Hepevirus in the new family Hepeviridae.43 HEV has been characterized as a nonenveloped 27–34 nm particle with a positive sense single-stranded RNA genome. Its genome is approximately 7.2 kb in length, and contains a short 5′ NTR, three ORFs (ORF1, ORF2, ORF3), 3′ NTR, and followed by a poly(A) tail.59–61 ORF1 is the largest among the three ORFs and encodes a nonstructural protein containing methyltransferase, helicase, and replicase domains. ORF2 encodes the major capsid protein of the virus, while ORF3 encodes a small protein that appears to serve as a viral regulatory protein rather than a structural protein.43,62,63 5.1.2.3 HEV Classification and Genetic Diversity Only a single known serotype of HEV is recognized to date. However, based on sequence analysis, several HEV isolates

Molecular Detection of Foodborne Pathogens

identified from different parts of the world have been classified into five genotypes.43,64,65 Of these, genotypes 1–4 are found in humans, genotypes 3 and 4 are restricted mainly in pigs, whereas genotype 5 has been isolated from chickens in the US.65 Data of global distribution of HEV in humans demonstrated that genotype 1 has been circulated in the developing countries in Asia and Africa, and is responsible mainly for waterborne outbreaks in these areas. Genotype 2 has been reported from outbreaks of HEV in Mexico and Nigeria. On the other hand, genotypes 3 and 4 are sometimes found to be associated with zoonotic infection and have been reported from sporadic cases of an acute HEV infection and/or domestic pigs in America, Europe, and Asia.43,47 5.1.2.4  Foodborne and Waterborne HEV HEV is spread mainly through the fecal-oral route via fecal contamination in water and food products. The growing data on the epidemiological studies of HEV indicated that HEV may be a zoonotic virus. Detections of HEV RNA from feces and anti-HEV from sera of swine have recently been documented.66–68 In addition, a number of studies reported the evidence of zoonotic foodborne transmission of HEV from animals to humans, such as, detection of HEV from pig livers sold in Japan and the US.57,69 Direct evidence for zoonotic foodborne transmission of HEV has also been confirmed in Japan where consumption of uncooked deer and wild-boar meat resulted in HEV infection in humans.70,71 In addition, waterborne outbreaks of HEV disease have been reported in association with contaminated water supply in several countries worldwide.72–76 5.1.2.5 Diagnosis of HEV For diagnosis of HEV infection, several methods are available. These include immune electron microscopy (IEM), enzyme immunoassay (EIA), immunochromatography (IC), RT-PCR, real-time PCR, and nucleic acid sequence analysis. Among these methods, RT-PCR and nucleic acid sequence analysis are widely used methods for the detection and genotype identification of foodborne HEV (food/water contaminations) and zoonotic HEV infections.77–80 These techniques have replaced the traditional immunological tests and become the gold standard for diagnosis of HEV infection. Recently, molecular techniques based on RT-PCR for the detection of viral genomic RNA have been developed for both epidemiological study and diagnosis of HEV infection. The most reliable marker for diagnosis of HEV infection is the presence of HEV RNA in serum or fecal samples. Therefore, several protocols of nested RT-PCR have been reported for direct detection of HEV from clinical specimens.77,81–88 The oligonucleotide primers used for the detection of HEV from clinical specimens are summarized in Table 5.2.

67

Hepatitis A and E Viruses

Table 5.2 Oligonucleotide Primers for the Detection and Sequencing of HEV From Clinical Specimens Primer Name

Region

Sequence 5′–3′

Position

Sense

HE361 HE364 HE366 HE363

ORF2/3 ORF2/3 ORF2/3 ORF2/3

First PCR First PCR Second PCR Second PCR

GCR GTG GTT TCT GGG GTG AC CTG GGM YTG GTC DCG CCA AG GYT GAT TCT CAG CCC TTC GC GYM TGG TCD CGC CAA GHG GA

5302–5321 5446–5465 5325–5344 5442–5461

+ – + –

ConsORF2-s1 ConsORF2-a1 ConsORF2-s2 ConsORF2-a2

ORF2 ORF2 ORF2 ORF2

First PCR First PCR Second PCR Second PCR

GAC AGA ATT RAT TTC GTC GGC TGG CTT GTT CRT GYT GGT TRT CAT AAT C GTY GTC TCR GCC AAT GGC GAG C GTT CRT GYT GGT TRT CAT AAT CCT G

6298–6321 6470–6494 6347–6368 6467–6491

+ – + –

E1 E5 E2 E4-1

ORF2 ORF2 ORF2 ORF2

First PCR First PCR Second PCR Second PCR

CTG TTT AAY CTT GCT GAC AC WGA RAG CCA AAG CAC ATC GAC AGA ATT GAT TTC GTC G TGY TGG TTR TCR TAA TC

6260–6279 6551–6568 6298–6316 6467–6486

+ – + –

F1 R1 F2 R2

ORF2 ORF2 ORF2 ORF2

First PCR First PCR Second PCR Second PCR

GCC GAG TAT GAC CAG TCC A ACA ACT CCC GAG TTT TAC CC AAT GTT GCG ACC GGC GCG C TAA GGC GCT GAA GCT CAG C

6577–6595 7107–7127 6649–6668 7079–7098

+ – + –

Forward (ex) Backward (ex) Forward (in) Backward (in)

ORF1 ORF1 ORF1 ORF1

First PCR First PCR Second PCR Second PCR

CAT GGT CGA GAA GGG CCA GG GCG GAA GTC ATA ACA GTG GG ATG ACT TTG CTG AGT TTG ACT CAT ATT CCA GAC AGT ATT CC

4089–4108 4631–4650 4403–4423 4601–4620

+ – + –

3156N 3157N 3158N 3159N

ORF2 ORF2 ORF2 ORF2

First PCR First PCR Second PCR Second PCR

AAT TAT GCY CAG TAY CGR GTT G CCC TTR TCY TGC TGM GCA TTC TC GTW ATG CTY TGC ATW CAT GGC T AGC CGA CGA AAT CAA TTC TGT C

5711–5732 6419–6441 5996–6017 6322–6343

+ – + –

PCR Product

Reference 77 77 77 77

164 bp 137 bp

82 82 82 82

197 bp 145 bp

88 88 88 88

309 bp 189 bp

84 84 84 84

551 bp 450 bp

86 86 86 86

562 bp 218 bp

85 85 85 85

731 bp 348 bp

5.2  Methods 5.2.1 Reagents and Equipments Reagents DEPC-treated water Ethanol Phosphate buffered saline (PBS), pH 7.4 Chloroform Isopropanol TRIzol® LS reagent QIAamp Viral RNA Mini Kit 5× First-Strand RT Buffer DTT Deoxynucleotide triphosphate (dNTP) Mix (2.5 and 10 mM) SuperScript III Reverse Transcriptase (200 units/µl) Random primer (1 µg/µl) RNase Inhibitor (40 units//µl) Mineral oil 5× Colorless GoTaq PCR buffer

Supplies and Equipments 0.2, 0.5, 1.5-ml tubes (RNase-free) Micropipetters and aerosol barrier filter tips (10, 100, 1000 µl) Disposable protective gloves Appropriate tube racks Suitable containers for ice Disposable plastic pipets Freezer –20, –80°C High speed microcentrifuge Class II biological safety cabinet Speed-vacuum Heat box Vortex mixer PCR thermal cycler Laboratory balances Agarose gel electrophoresis apparatus Timers (Continued)

68

Molecular Detection of Foodborne Pathogens

Reagents

Supplies and Equipments

Specific primers for HAV and HEV GoTaq DNA polymerase (5 units/µl) TAE (Tris-acetate-EDTA) buffer Agaro Loading dye 100 bp DNA ladder marker Ethidium bromide Wizard SV Gel and PCR Clean-Up System or QIAquick PCR purification kit BigDye Terminator v3.1 Cycle Sequencing Kit

5.2.2 Sample Collection and Preparation 5.2.2.1 HAV Specimen Collection and RNA Extraction Although feces, serum, saliva, and liver suspension can be used for diagnosis of foodborne HAV infection, the specimens of choice are feces and serum. The fecal specimen should be collected within 1 week after the onset of dark urine and stored at –20°C until tested.89 The 10% stool suspension is prepared in phosphate-buffered saline (pH 7.4) and clarified by centrifugation at 4,800 × g for 15 min to eliminate larger debris. Serum sample should be collected during the first month after presentation of clinical signs or symptoms (jaundice, dark urine, or the level of serum ALT/AST are higher than 20 UI/L) The obtained serum should be stored at –20°C or below until use for RNA extraction. For the TRIzol® LS extraction method, 100 µl of serum or 10% (w/v) fecal supernatant is mixed with 800 µl of TRIzol® LS reagent and 200 µl of chloroform. After centrifugation at 8,000 × g for 20 min at 4°C, the upper phase is precipitated with 600 µl of isopropanol at –20°C for 18–24 h. Then, follows by centrifugation, and the pellet is washed with 70% ethanol, air dry, and finally resuspended in 10 µl of DEPCtreated water. For QIAamp Viral RNA Mini Kit, the RNA is extracted from the fecal supernatant or serum according to the manufacturer’s instructions. Briefly, 140 µl of 10% (w/v) fecal supernatant or serum is mixed with 560 µl of AVL viral lysis buffer. The mixture is incubated at room temperature for 10 min and 560 µl of ethanol is added. Then, the mixture is applied onto the spin column, centrifuge at 6000 × g for 1 min, and 500 µl of AW1 buffer is added. The column is centrifuged at 6,000 × g for 1 min to remove unbound materials, and washed by addition of 500 µl of AW2 buffer. Then, the column is centrifuged at full-speed for 3 min, and placed into a new 1.5-ml microcentrifuge tube. Finally, 60 µl of AVE buffer is added directly onto the column to elute HAV genomic RNA. After incubation at room temperature for 1 min, the column is centrifuged at 6,000 × g for 1 min. The obtained RNA is used as a template for PCR amplification. The genomic RNA can be stored at –80°C until nested RT-PCR assay is performed. 5.2.2.2 HEV Specimen Collection and RNA Extraction The specimens of choice for the diagnosis of HEV infection are serum and stool samples. Specimens should be collected

Ultraviolet transilluminator Syringe and needle Sterile Pasteur piptes DNA sequencer (ABI 3100 Applied Biosystems)

in an early acute phase of the disease and stored at –20°C until tested. Long-term storage should be kept at –70°C. Several methods have been used for the extraction of HEV RNA from clinical specimens prior to RT-PCR. These include the use of glass powder RNA extraction, antigencapture method by the use of anti-HEV antibody to capture the virus particle, microspin columns for HEV RNA extraction, and conventional RNA separation by using GTCphenol-chloroform extraction method.90–92 Recently, highly sensitive and reproducible RNA extraction kits, which are commercially available, have been introduced and widely used for HEV genomic RNA isolation, including Trizol LS® Reagent (Invitrogen Life Technology) and QIAamp Viral RNA Mini Kit (Qiagen).77,80–83 The details of specimen collection and RNA extraction methods are similar to those of the protocols described above for HAV detection.

5.2.3  Detection Procedures 5.2.3.1 Detection of HAV (i) Reverse-transcription PCR (RT-PCR): The following RT-PCR protocol has been modified from previously published reports.24,26,27 The RT reaction is carried out in a total volume of 15 µl. Note: Vortex and spin down all microcentrifuge tubes of RT-PCR components before use. (1) Prepare the reverse transcription mix (RT-mix) as follows (volumes indicated as per specimen): Component DEPC-treated water 5× First-strand buffer 0.1 M DTT dNTP mix (10 mM) SuperScript III reverse transcriptase Random primer (hexa-deoxyribonucleotide mixture; 1 µg/µl) RNase inhibitor Total



Volume (µl) 3.3 3.0 0.8 0.8 0.8 0.8 0.5 10.0

(2) Add 10 µl RT-mix to each 0.5 ml tube followed by addition of 5.0 µl of individual RNA sample. (3) Spin the tubes briefly to ensure that no reagent droplets remain on the side of the tubes. (4) Overlay the reaction mixture with one drop of mineral oil to prevent evaporation during the RT process.

69

Hepatitis A and E Viruses







(5) Transfer the reaction tubes to a 50°C heat box for 1 h, and then increase temperature to 95°C for 5 min. (6) Rapidly chill the tubes on ice for 5 min. (7) The cDNA can be used directly in the PCR, or stored at –20°C for future use. (ii) Nested PCR for HAV detection: For nested PCR amplification, two different PCR reactions (first and second PCR) are performed separately in a total volume of 25 µl. The cDNA is amplified for the detection of HAV genome. A forward primer BR-5 is used in combination with a reverse primer BR-9 (Table 5.1) to specifically generate a PCR amplicon of 360 bp.24 (1) Remove all aliquots of PCR reagents (DEPCtreated water, 5 × Colorless GoTaq PCR buffer, 2.5 mM dNTP mix, primers, GoTaq DNA polymerase) from the –20°C freezer. Vortex and spin down all reagents before opening the tubes. (2) Prepare the first PCR reaction mix as follows (volumes indicated as per specimen) : Component

Volume (µl)

DEPC-treated water

13.9 5.0

5 × Colorless GoTaq PCR buffer (containing MgCl2) dNTP mix (2.5 mM) Primer BR-5 (20 pmol/µl)

2.0 0.5

Primer BR-9 (20 pmol/µl)

0.5 0.1

GoTaq DNA polymerase (5 units/µl) Total





22.0

(3) Add 22.0 µl of first PCR mix and 3.0 µl of cDNA into a 0.2-ml PCR tube. (4) Turn on a thermocycler and preheat the block to 93°C. (5) Spin the sample tubes in a microcentrifuge at 14,000 × g for 30 sec at room temperature. (6) Place the sample tubes in the thermal cycler for 30 cycles of 93°C for 30 sec, 55°C for 30 sec, 72°C for 50 sec; and one cycle of 72°C for 10 min. (7) Following amplification, pulse-spin the reaction tubes to pull down the condensation droplets at the inner wall of the tubes. The samples are now ready for the second PCR or stored frozen at –20°C. (8) The second PCR is conducted using the first PCR product as a template. Prepare the second PCR reaction mix as follows (volumes indicated as per specimen): Component DEPC-treated water 5 × Colorless GoTaq PCR buffer (containing MgCl2) dNTP mix (2.5 mM)



(9) Add 23.0 µl of second PCR mix and 2.0 µl of first PCR product to a 0.2-ml PCR tube. (10) Then, the PCR is carried out under the same thermal cycling conditions as the first amplification. The expected size of the second PCR product is 234 bp in length. (11) Analyze the PCR products by agarose gel electrophoresis through 2% agarose gel in TAE buffer at 100 volts for 30 min. (12) The gel is stained with ethidium bromide and then visualized under ultraviolet light source. Note: negative control is also concurrently included along with the test samples in order to monitor any possible contamination that might be occur during the PCR procedure. (iii) HAV sequence analysis and genotype identification: To characterize HAV genotypes, a phylogenetic analysis should be carried out by direct sequencing of the nucleotide sequence of VP1/P2A junction. A key step in all sequencing methods is the careful preparation of the template with adequate concentration and purity. Prior to sequencing reaction, the unincorporated nucleotide and primers remaining after the PCR amplification must be removed. This step can be done simply by using a commercial spin column kits (Wizard SV Gel and PCR Clean-Up System; Promega, Madison, WI or QIAquick PCR purification kit; Qiagen). The amount of purified PCR product is once more estimated by electrophoresis through 2% agarose gel in parallel with standard DNA and the purified product is used as a template in cycle sequencing reaction. Nucleotide sequencing is performed using the BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA) according to the manufacturer protocol. (1) For each sequencing reaction, mix the following reagents in a labeled tube Component Terminator ready reaction mix (v3.1) 5 × Sequencing buffer Purified PCR product (~100 ng) Sequencing primer (BR-6; 5 pmol/µl) DEPC-treated water Total



Volume (µl) 14.9 5.0

Primer RJ-3 (20 pmol/µl)

2.0 0.5

Primer BR-6 (20 pmol/µl)

0.5

GoTaq DNA polymerase (5 units/µl) Total

0.1 23.0





Volume (µl) 4.0 4.0 1.0 1.0 10.0 20.0

(2) Spin the samples tubes in a microcentrifuge at 14,000 × g for 30 sec at room temperature. (3) Place the sample tubes in a thermocycler for a total of 25 cycles of 96°C for 10 sec, 50°C for 5 sec, and 60°C for 4 min. (4) Then, the DNA sequencing product is purified by ethanol-EDTA precipitation, washed with 70% ethanol, and dry the pellet in a vacuum centrifuge. (5) Finally, analyze the nucleotide sequence of the DNA product using an automated DNA sequencer ABI 3100 Applied Biosystems.

70

Molecular Detection of Foodborne Pathogens

Note: having obtained good-quality sequence data, it is necessary to compare the sequence data with other published sequences. Therefore, the HAV nucleotide sequences of VP1/P2A junction should be compared with those of the reference strains available in the NCBI (National Center for Biotechnology Information) GenBank database, EMBL (The European Molecular Biology Laboratory), or DDBJ (DNA Data Bank of Japan). Phylogenetic and molecular evolutionary analyses are conducted using MEGA version 4.93 The authors strongly recommend that the new sequences obtained should be submitted for deposition in one of the databases mentioned above, in order to make them available to the scientific community. Most journals do not accept manuscripts before deposition of the sequence data in one of those databases. 5.2.3.2 Detection of HEV (i) RT-PCR: The protocols that are routinely used for the detection and genotyping of HEV from clinical specimens have been modified from References 77 and 81. RNA is reverse-transcribed in the presence of random primers and the RT is carried out under the same conditions as those described above for HAV detection. (ii) Nested PCR for HEV detection: To detect the HEV RNA, a part of ORF2/3 is amplified by nested PCR. The oligonucleotide primers for the detection of HEV are described previously by Inoue et al. (Table 5.2).77 The reaction mixture for the first round amplification is similar to that of HAV except for the primers used. (1) Remove all aliquots of PCR reagents (DEPCtreated water, 5 × Colorless GoTaq PCR buffer, 2.5 mM dNTP mix, primers, GoTaq DNA polymerase) from the –20°C freezer. Vortex and spin down all reagents before open the tubes. (2) Prepare the first PCR reaction mix as follow (volumes indicated as per specimen): Component DEPC-treated water 5 × Colorless GoTaq PCR buffer (containing MgCl2) dNTP mix (2.5 mM)



13.9 5.0

Primer HE361 (20 pmol/µl)

2.0 0.5

Primer HE364 (20 pmol/µl)

0.5

GoTaq DNA polymerase (5 units/µl) Total



Volume (µl)

0.1 22.0

(3) Add 22.0 µl of first PCR mix and 3.0 µl of cDNA into a 0.2-ml PCR tube. (4) Turn on a thermal cycler and preheat the block to 94°C. (5) Spin the sample tubes in a microcentrifuge at 14,000 × g for 30 sec at room temperature.







(6) Place the sample tubes in the thermocycler for a total of 35 cycles of 94°C for 30 sec, 58°C for 30 sec, and 72°C for 30 sec; and one cycle of 72°C for 10 min. (7) Following amplification, pulse-spin the reaction tubes to pull down all the solution that may attach to the inner wall of the tubes. The samples are ready for second PCR or stored frozen at –20°C. (8) Then, nested PCR is performed in a total volume of 25 µl with 2 µl of amplification product from the first round PCR. Prepare the second PCR reaction mix as follows (volumes indicated as per specimen): Component DEPC-treated water 5 × Colorless GoTaq PCR buffer (containing MgCl2) dNTP mix (2.5 mM)

Volume (µl) 14.9 5.0

Primer HE366 (20 pmol/µl)

2.0 0.5

Primer HE363 (20 pmol/µl)

0.5

GoTaq DNA polymerase (5 units/µl) Total

0.1 23.0



(9) Add 23.0 µl of second PCR mix and 2.0 µl of first PCR product to a 0.2-ml PCR tube. (10) Then, the nested PCR is carried out under the same thermal cycling conditions as the first amplification. The expected sizes of the amplification products are 164 bp and 137 bp for the first and second round PCR, respectively. (11) The PCR product sizes are determined by electrophoresis through 3% agarose gel in TAE buffer at 100 volts for 30 min. (12) The gel is stained with ethidium bromide and then visualized under ultraviolet light source. Note: negative control is also concurrently included along with the test samples in order to monitor any possible contamination that might occur during the PCR procedure. (iii) HEV sequence analysis and genotype identification: For the detection of genetic variation and genotype identification of HEV, genome sequence and phylogenetic analyses should be performed at least one part of the gene by direct sequencing of PCR products. HEV PCR product purification and sequencing protocol are similar to those described above for the HAV sequence analysis except for sequencing primer HE363 is used. (1) For each sequencing reaction, mix the following reagents in a labeled tube Component Terminator ready reaction mix (v3.1) 5 × Sequencing buffer Purified PCR product (~100 ng) Sequencing primer (HE363; 5 pmol/µl) DEPC-treated water Total

Volume (µl) 4.0 4.0 1.0 1.0 10.0 20.0

71

Hepatitis A and E Viruses





(2) Spin the samples tubes in a microcentrifuge at 14,000 × g for 30 sec at room temperature. (3) Place the sample tubes in a thermocycler for 25 cycles of 96°C for 10 sec, 50°C for 5 sec, and 60°C for 4 min. (4) Then, the DNA sequencing product is purified by ethanol-EDTA precipitation, washed with 70% ethanol, and the pellet is dried in a vacuum centrifuge. (5) Finally, the nucleotide sequence of the DNA product is analyzed using an automated DNA sequencer ABI 3100 Applied Biosystems.

5.3 Conclusions and Future Perspectives Acute viral hepatitis is an important disease caused by a variety of hepatitis viruses. Among them, HAV, and HEV contribute a significant role on the disease. Both are enteric viruses and transmitted by fecal-oral contact or consumption of contaminated food or water. HAV is responsible for acute viral hepatitis, which represents a significant public health problem in many countries due to the persistent circulation of the viruses in the environment. HEV epidemics have also been reported in some developing countries and are associated frequently with contaminated drinking water. In addition, it causes substantial morbidity and mortality, particularly among pregnant women. Recently, the determination and analysis of molecular biomarkers (e.g., nucleic acid sequencing or phylogenetic analysis) have been used to detect and to determine the relatedness of the viruses circulating worldwide. Molecular epidemiology has become a particularly powerful tool for investigation of foodborne and waterborne outbreaks of the diseases. In addition, the availability of a large and growing database of HAV and HEV sequences has allowed the identification of the source of virus contamination and may help to accelerate the public health concern and immediate response to the outbreaks. Although single round PCR is the most familiar molecular assay for the detection and genotyping of pathogens in clinical microbiology, the multiplex PCR assay is also getting much attention and has been developed and widely used to serve these purposes in several viruses. However, multiplex PCR has some inherited limitations, particularly the sensitivity of the method. For future perspective, a more highly sensitive, reliable, and inexpensive method needs to be developed for the detection and genotyping of pathogens. Most recently, DNA microarray technology has been developed and widely applicable in several aspects of infectious diseases, particularly for the detection and identification of infecting pathogens. In some cases, the detection limit of microarray was found to be as little as 10 fg of pathogenic DNA, which is far below the detection limits of other existing technologies. Conceivably, DNA microarray is an ideal tool that needs to be developed for the detection and identification, as well as genotyping and characterizing of HAV and HEV in the future.

References











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72 23. Apaire-Marchais, V. et al. Development of RT-semi-nested PCR for detection of hepatitis A virus in stool in epidemic conditions. Mol. Cell. Probes, 8, 117, 1994. 24. Bruisten, S.M. et al. Molecular epidemiology of hepatitis A virus in Amsterdam, the Netherlands. J. Med. Virol., 63, 88, 2001. 25. Chironna, M. et al. Genetic analysis of HAV strains recovered from patients with acute hepatitis from Southern Italy. J. Med. Virol., 70, 343, 2003. 26. Poovorawan, Y. et al. Clinical features and molecular characterization of hepatitis A virus outbreak in a child care center in Thailand. J. Clin. Virol., 32, 24, 2005. 27. Rodrigues, L. et al. Molecular epidemiology of hepatitis A virus in a group of Portuguese citizens living in Lisbon area. J. Med. Virol., 79, 483, 2007. 28. de Paula, V.S. et al. Detection of hepatitis A virus RNA in serum during the window period of infection. J. Clin. Virol., 29, 254, 2004. 29. Costa-Mattioli, M. et al. Molecular evolution of hepatitis A virus: a new classification based on the complete VP1 protein. J. Virol., 76, 9516, 2002. 30. Endo, K. et al. Analysis of the full-length genome of a subgenotype IIIB hepatitis A virus isolate: primers for broadly reactive PCR and genotypic analysis. J. Med. Virol., 79, 8, 2007. 31. Amado, L.A. et al. Comparison between serum and saliva for the detection of hepatitis A virus RNA. J. Virol. Methods, 148, 74, 2008. 32. Goswami, B.B., Burkhardt, W., and Cebula, T.A. Identification of genetic variants of hepatitis A virus. J. Virol. Methods, 65, 95, 1997. 33. Buti, M. et al. Assessment of the PCR-Southern blot technique for the analysis of viremia in patients with acute hepatitis A. Gastroenterol. Hepatol., 24, 1, 2001. 34. Shan, X.C., Wolffs, P., and Griffiths, M.W. Rapid and quantitative detection of hepatitis A virus from green onion and strawberry rinses by use of real-time reverse transcriptionPCR. Appl. Environ. Microbiol., 71, 5624, 2005. 35. Yoneyama, T. et al. Rapid and real-time detection of ­hepatitis A  virus by reverse transcription loop-mediated isothermal amplification assay. J. Virol. Methods, 145, 162, 2007. 36. Papafragkou, E. et al. Rapid and sensitive detection of hepatitis A virus in representative food matrices. J. Virol. Methods, 147, 177, 2008. 37. Chomczynski, P., and Sacchi, N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem., 162, 156, 1987. 38. Boom, R. et al. Rapid and simple method for purification of nucleic acids. J. Clin. Microbiol., 28, 495, 1990. 39. Deng, M.Y., Day, S.P., and Cliver, D.O. Detection of hepatitis A virus in environmental samples by antigen-capture PCR. Appl. Environ. Microbiol., 60, 1927, 1994. 40. Jothikumar, N., Cliver, D.O., and Mariam, T.W. Immuno­ magnetic capture PCR for rapid concentration and detection of hepatitis A virus from environmental samples. Appl. Environ. Microbiol., 64, 504, 1998. 41. de Paula, V.S. et al. Comparison of four extraction methods to detect hepatitis A virus RNA in serum and stool samples. Braz. J. Infect. Dis., 7, 135, 2003. 42. Phan, T.G. et al. A novel RT-multiplex PCR for enteroviruses, hepatitis A and E viruses and influenza A virus among infants and children with diarrhea in Vietnam. Arch. Virol., 150, 1175, 2005. 43. Emerson, S.U., and Purcell R.H. Hepatitis E Virus. In: Knipe, D.M, Howley, P.M., (ed.), Fields Virology, 5th ed. pp. 3047– 3058. Lippintcott Williams & Wilkins, Philadelphia, PA, 2007.

Molecular Detection of Foodborne Pathogens 44. Aggarwal, R., and Krawczynski, K. Hepatitis E: an overview and recent advances in clinical and laboratory research. J. Gastroenterol. Hepatol., 15, 9, 2000. 45. Emerson, S.U., and Purcell, R.H. Hepatitis E virus. Rev. Med. Virol., 13, 145, 2003. 46. Krawczynski, K. Hepatitis E. Hepatology, 17, 932, 1993. 47. Mushahwar, I.K. Hepatitis E virus: molecular virology, clinical features, diagnosis, transmission, epidemiology, and prevention. J. Med. Virol., 80, 646, 2008. 48. Jilani, N. et al. Hepatitis E virus infection and fulminant hepatic failure during pregnancy. J. Gastroenterol. Hepatol., 22, 676, 2007. 49. Tsega, E. et al. Acute sporadic viral hepatitis in Ethiopia: causes, risk factors, and effects on pregnancy. Clin. Infect. Dis., 14, 961, 1992. 50. Khuroo, M.S. and Kamili, S. Aetiology, clinical course and outcome of sporadic acute viral hepatitis in pregnancy. J. Viral. Hepatol., 10, 61, 2003. 51. Chauhan, A. et al. Hepatitis E virus transmission to a volunteer. Lancet, 16, 149, 1993. 52. Tsarev, S.A. et al. Characterization of a prototype strain of hepatitis E virus. Proc. Natl. Acad. Sci. USA, 15, 559, 1992. 53. Balayan, M.S. et al. Evidence for a virus in non-A, non-B hepatitis transmitted via the fecal-oral route. Intervirology. 20, 23, 1983. 54. Purcell, R.H. and Emerson, S.U. Animal models of hepatitis A and E. ILAR. J., 42, 161, 2001. 55. Okamoto, H., Takahashi, M., and Nishizawa, T. Features of hepatitis E virus infection in Japan. Intern. Med., 42, 1065, 2003. 56. Divizia, M. et al. Waterborne gastroenteritis outbreak in Albania. Water. Sci. Technol., 50, 57, 2004. 57. Yazaki, Y. et al. Sporadic acute or fulminant hepatitis E in Hokkaido, Japan, may be food-borne, as suggested by the presence of hepatitis E virus in pig liver as food. J. Gen. Virol., 84, 2351, 2003. 58. Emerson, S.U., Arankalle, V.A., and Purcell, R.H. Thermal stability of hepatitis E virus. J. Infect. Dis., 192, 930, 2005. 59. Reyes, G.R. et al. Isolation of a cDNA from the virus responsible for enterically transmitted non-A, non-B hepatitis. Science, 247, 1335, 1990. 60. Tam, A.W. et al. Hepatitis E virus (HEV): molecular cloning and sequencing of the full-length viral genome. Virology, 185, 120, 1991. 61. Wang, Y. et al. The complete sequence of hepatitis E virus genotype 4 reveals an alternative strategy for translation of open reading frames 2 and 3. J. Gen. Virol., 81, 1675, 2000. 62. Koonin, E.V. et al. Computer-assisted assignment of functional domains in the nonstructural polyprotein of hepatitis E virus: delineation of an additional group of positive-strand RNA plant and animal viruses. Proc. Natl. Acad. Sci. USA, 89, 8259, 1992. 63. Surjit, M., Jameel, S., and Lal, S.K. The ORF2 protein of hepatitis E virus binds the 5' region of viral RNA. J. Virol., 78, 320, 2004. 64. Emerson, S.U., and Purcell, R.H. Hepatitis E. Pediatr. Infect. Dis. J., 26, 1147, 2007. 65. Purcell, R.H., and Emerson, S.U. Hepatitis E: an emerging awareness of an old disease. J. Hepatol., 48, 494, 2008. 66. Clayson, E.T. et al. Detection of hepatitis E virus infections among domestic swine in the Kathmandu Valley of Nepal. Am. J. Trop. Med. Hyg., 53, 228, 1995. 67. Arankalle, V.A. et al. Human and swine hepatitis E viruses from Western India belong to different genotypes. J. Hepatol., 36, 417, 2002.

Hepatitis A and E Viruses 68. Fernández-Barredo, S. et al. Prevalence and genetic ­characterization of hepatitis E virus in paired samples of feces and serum from naturally infected pigs. Can. J. Vet. Res., 71, 236, 2007. 69. Feagins, A.R. et al. Detection and characterization of infectious Hepatitis E virus from commercial pig livers sold in local grocery stores in the USA. J. Gen. Virol., 88, 912, 2007. 70. Tei, S. et al. Consumption of uncooked deer meat as a risk factor for hepatitis E virus infection: an age- and sex-matched case-control study. J. Med. Virol., 74, 67, 2004. 71. Masuda, J. et al. Acute hepatitis E of a man who consumed wild boar meat prior to the onset of illness in Nagasaki, Japan. Hepatol. Res., 31, 178, 2005. 72. Rab, M.A. et al. Water-borne hepatitis E virus epidemic in Islamabad, Pakistan: a common source outbreak traced to the malfunction of a modern water treatment plant. Am. J. Trop. Med. Hyg., 57, 151, 1997. 73. Sarguna, P., Rao, A., and Sudha Ramana, K.N. Outbreak of acute viral hepatitis due to hepatitis E virus in Hyderabad, India. J. Med. Microbiol., 25, 378, 2007. 74. Maila, H.T., Bowyer, S.M., and Swanepoel, R. Identification of a new strain of hepatitis E virus from an outbreak in Namibia in 1995. J. Gen. Virol., 85, 89, 2004. 75. Bile, K. et al. Contrasting roles of rivers and wells as sources of drinking water on attack and fatality rates in a hepatitis E epidemic in Somalia. Am. J. Trop. Med. Hyg., 51, 466, 1994. 76. Panda, S.K., Thakral, D., and Rehman, S. Hepatitis E virus. Rev. Med. Virol., 17, 151, 2007. 77. Inoue, J. et al. Development and validation of an improved RT-PCR assay with nested universal primers for detection of hepatitis E virus strains with significant sequence divergence. J. Virol. Methods, 137, 325, 2006. 78. Jothikumar, N. et al. Detection of hepatitis E virus in raw and treated wastewater with the polymerase chain reaction. Appl. Environ. Microbiol., 59, 2558, 1993. 79. Huang, F.F. et al. Detection by reverse transcription-PCR and genetic characterization of field isolates of swine hepatitis E virus from pigs in different geographic regions of the United States. J. Clin. Microbiol., 40, 1326, 2002. 80. Garkavenko, O. et al. Detection and characterisation of swine hepatitis E virus in New Zealand. J. Med. Virol., 65, 525, 2001.

73 81. Takahashi, M. et al. Prolonged fecal shedding of hepatitis E virus (HEV) during sporadic acute hepatitis E: evaluation of infectivity of HEV in fecal specimens in a cell culture system. J. Clin. Microbiol., 45, 3671, 2007. 82. Zhao, C. et al. Comparison of real-time fluorescent RT-PCR and conventional RT-PCR for the detection of hepatitis E virus genotypes prevalent in China. J. Med. Virol., 79, 1966, 2007. 83. Mizuo, H. et al. Polyphyletic strains of hepatitis E virus are responsible for sporadic cases of acute hepatitis in Japan. J. Clin. Microbiol., 40, 3209, 2002. 84. He, J. Molecular detection and sequence analysis of a new hepatitis E virus isolate from Pakistan. J. Viral. Hepatol., 13, 840, 2006. 85. Cooper, K. et al. Identification of genotype 3 hepatitis E virus (HEV) in serum and fecal samples from pigs in Thailand and Mexico, where genotype 1 and 2 HEV strains are prevalent in the respective human populations. J. Clin. Microbiol., 43, 1684, 2005. 86. Zhao, Z.Y. et al. Detection of hepatitis E virus RNA in sera of patients with hepatitis E by polymerase chain reaction. Hepatobiliary Pancreat. Dis. Int., 6, 38, 2007. 87. Aggarwal, R. et al. Genetic variability of hepatitis E virus within and between three epidemics in India. Virus. Res., 59, 35, 1999. 88. Zhang, F. et al. Detection of HEV antigen as a novel marker for the diagnosis of hepatitis E. J. Med. Virol., 78, 1441, 2006. 89. Coulepis, A.G. et al. Detection of hepatitis A virus in the feces of patients with naturally acquired infections. J. Infect. Dis., 141, 151, 1980. 90. McCaustland, K.A. et al. Application of two RNA extraction methods prior to amplification of hepatitis E virus nucleic acid by the polymerase chain reaction. J. Virol. Methods, 35, 331, 1991. 91. Ray, R. et al. Hepatitis E virus genome in stools of hepatitis patients during large epidemic in north India. Lancet, 338, 783, 1991. 92. Aggarwal, R., and McCaustland, K.A. Hepatitis E virus RNA detection in serum and feces specimens with the use of microspin columns. J. Virol. Methods, 74, 209, 1998. 93. Tamura, K. et al. MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Mol. Biol. Evol., 24, 1596, 2007.

6 Noroviruses Anna Charlotte Schultz

Technical University of Denmark (DTU)

Jan Vinjé

Center for Disease Control (CDC)

Birgit Nørrung

University of Copenhagen

Contents 6.1 Introduction......................................................................................................................................................................... 75 6.1.1 Background and Classification................................................................................................................................ 75 6.1.2 Biology..................................................................................................................................................................... 76 6.1.3 Pathogenesis............................................................................................................................................................. 76 6.1.4 Epidemiology........................................................................................................................................................... 76 6.1.5 Diagnosis.................................................................................................................................................................. 77 6.1.5.1 Issues Concerning Sample Processing...................................................................................................... 77 6.1.5.2 Issues Concerning Amplification, Detection, and Quality Assurance (QA)............................................. 80 6.2 Methods............................................................................................................................................................................... 82 6.2.1 Reagents and Equipment.......................................................................................................................................... 82 6.2.2 Sample Preparation.................................................................................................................................................. 82 6.2.3 Detection of Noroviruses by Real-time RT-PCR.................................................................................................... 85 6.3 Conclusion and Future Perspectives.................................................................................................................................... 85 Acknowledgments........................................................................................................................................................................ 85 References.................................................................................................................................................................................... 85

6.1  Introduction

6.1.1  Background and Classification

Noroviruses (NoVs) have emerged as a major cause of foodborne illness causing an estimated 30–50% of all foodborne outbreaks in the U.S. Because these viruses are highly contagious, have a low infectious dose, and are persistent in the environment, they are classified as group B biodefense pathogens. The source of most foodborne outbreaks is contaminated bivalve shellfish, which are often eaten raw, or infected food handlers. Foods that have been implicated in outbreaks include ready-to-eat (RTE) foods such as salads, celery, melon, sandwiches, lettuce, cold cooked ham, and fresh or frozen raspberries. Detection of NoV in food is limited to molecular methods as the virus cannot be grown in routine cell culture. This chapter describes the history, classification, and biology of NoVs and provides an overview of the published methods for the molecular detection in various food items. Detailed laboratory protocols for the rapid concentration, extraction, and real-time reverse transcriptase-polymerase chain reaction (RT-PCR) detection in bivalve molluscan shellfish and raspberries are provided.

Although outbreaks of nonbacterial gastroenteritis have been recognized as early as the 1930s, a causative viral agent was not discovered until 1972 when 27 nm sized virus particles were detected by immune electron microscopy (EM) in stool samples of human volunteers who had been fed with a bacterial-free filtrate from an individual that was involved in an outbreak of acute gastroenteritis in an elementary school in Norwalk, Ohio in 1968.63 The virus was named Norwalk virus (NV) and this virus has been established as the prototype virus for the group of viruses now called NoVs, but in the literature also referred to as “Norwalk-like viruses” or small round structured viruses (SRSVs) based on the characteristic surface morphology of the virus particles. The classification of NoVs as members of the family Caliciviridae was established after the cloning and sequencing of the complete genome of NV58 and Southampton virus.72 To date, these viruses are grouped into a separate genus Norovirus. Other genera include Sapovirus, Lagovirus, and Vesivirus41,100 and a new fifth genus is pending. Below the genus level naming of NoVs has not been agreed upon and therefore NoV strains 75

76

are often named after the geographic location where the outbreaks first occurred such as Hawaii virus, Snow mountain virus, Bristol virus, and Minerva virus. A more formal naming system similar to influenza has been proposed with: species infected/virus genus/genogroup/virus name/strain designation/year of detection/country.4

6.1.2  Biology NoVs are a group of 27–35 nm nonenveloped viruses with an icosahedral capsid enclosing a single stranded positive sense RNA genome of approximately 7.5–7.7 nucleotides (nt). The NoV genome contains three open reading frames (ORFs) with ORF1 at the 5′-end of the genome which encodes a polyprotein of six nonstructural proteins including p48, NTPase, p22, VPg (binds to 5′ end to initiate translation), 3CLpro (protease), and RdRp (RNA dependent RNA polymerase). ORF2 and ORF3 code for two structural proteins: a major (VP1) and a minor (VP2) capsid protein.40 NoVs are a genetically diverse group of viruses which are currently classified into five genetic groups or genogroups.40,148 Strains that infect humans belong to genogroups I, II, and IV.148 Genogroup II includes also strains that have been detected uniquely in swine whereas viruses belonging to genogroup III and V infect cattle or mice, respectively. These five genogroups can be further divided into genotypes or genoclusters (eight in genogroup I, 19 in genogroup II, two in genogroup III, one in genogroup IV, and one in genogroup V).148 Subgenomic RNA, rather than genomic RNA, is used as the template for the production of the capsid proteins.8 Additionally, the 5′ end of the subgenomic and genomic RNAs contains a highly conserved nucleotide motif up-stream from the first AUG start codon and is therefore thought to be a site for the initiation of transcription.67 Using cryo-immune EM the structure of the NV has been resolved.111 Virus particles are assembled from 90 dimers of VP1 and a few VP2 molecules and form a T=3 icosahedral capsid.111 The exact role of VP2 is not completely known although this protein may be important in the stability of the overall viral capsid (capsid–RNA complex).8 VP1 folds into a shell domain (S) and a protruding domain (P) linked by an amino acid hinge.111 The relatively conserved S domain is essential for the formation of the icosahedral structure of the virion, while the P domain stabilizes the capsid by a dimeric interaction and forms the characteristic protrusions on the virion.46 The P domain is further subdivided into the P1 and P2 subdomains with the P2 subdomain containing both immune and cellular recognition sites.16,89,133

6.1.3  Pathogenesis After an incubation period between 24 and 48 hours but sometimes as short as 15 hours,38 clinical symptoms include acute projectile vomiting, nonbloody diarrhea, abdominal cramps, nausea, chills, myalgia, and headache.64 Fever is rarely reported. In human volunteers, NV has been shown to

Molecular Detection of Foodborne Pathogens

affect epithelial cells of the small intestine resulting in broadening and flattening of the villi. NoVs do not appear to infect the cells of the large intestine.29,120,127 During acute illness, the inflammation of the mucosa of the small intestine reduces the adsorptive capacity of the villi and produce intestinal fluid contributing to diarrhea while vomiting is likely due to delayed gastric emptying and disrupted motility.101 However, the precise mechanism of virus-induced diarrhea and vomiting is currently unknown.135 The disease in healthy individuals is usually mild and resolved after 1–3 days.64,65 However, severe illness has been reported in young children, the elderly, cardiovascular or renal disease patients,98 transplant recipients, and other immunosuppressed patients,55 in which case fluid replacement or additional supportive care may be needed.145 Rates of asymptomatic virus shedding of 32% have been reported in both clinical outbreak studies and studies in human volunteers experimentally infected with NV.38

6.1.4 Epidemiology NoVs are transmitted primarily through the fecal-oral route, either directly from person-to-person via contaminated hands, or indirectly via contaminated food or water. The virus can also persist for several days to weeks on surfaces and fomites which can act as a source of continuing infection during outbreaks. Importantly, airborne droplets of vomitus are infectious and have been strongly implicated in contaminating the environment and in direct transmission of NoV within semiclosed communities such as cruise ships, hospitals, the military and long-term care facilities.97 Human NoVs are estimated to be responsible for up to 95% of nonbacterial epidemic gastroenteritis worldwide in humans of all age groups.40 Recent studies indicate that NoV disease is also associated with a significant number of sporadic cases of gastroenteritis.42,56,94 Most outbreaks throughout the world are caused by strains that belong to GII, whereas <5% of the outbreaks are caused by GI and GIV outbreaks are very rare. In most years, viruses of different genotypes cocirculate but in 1996, 2002, and 2006, viruses of genetic cluster GII.4 were associated with the majority of the outbreaks globally. Molecular analysis of these GII.4 strains demonstrated that older GII.4 strains are being replaced by new GII.4 subtypes every 2–3 years coinciding with an increase in the number of outbreaks. These findings are somewhat similar to the evolution of influenza viruses and it has been hypothesized that herd immunity may contribute to a proposed epochal evolution model for GII.4 NoVs88,126 although there are other possible explanations that could describe these observations (e.g., natural fluctuation). NoVs bind to human histoblood group antigens (HBGAs),52,53,57,87,96,131,132 which are complex carbohydrates present on the surfaces of red blood cells as well as on the mucosal epithelium and in body fluids with important biological functions.95,112 The recognition of HBGAs by the capsid protein of NoVs appear to be highly strain specific,

Noroviruses

and eight distinct receptor-binding patterns have been iden­ tified.47,48,54 The binding of NV and other NoV genotypes to certain HBGA molecules has also been strongly linked with clinical infection.57,87,113 Recent studies have shown that HBGAs and in particular secretor status controlled by the 1,2-fucosyltransferase FUT2 gene determine susceptibility to NoV infections, with nonsecretors (FUT2−/−), representing 20% of Europeans, being highly resistant to symptomatic infections with major strains of NoV such as GII.4.73,134 Due to the different HBGA specificities of the capsid protein from distinct strains, a host–pathogen coevolution driven by carbohydrate–protein interactions has been suggested. For a comprehensive overview on HBGAs and susceptibility to NoV infection, the reader is referred to a review of LePendu et al.83 NoV outbreaks are often linked to food handlers who infect foods that are eaten raw or not further processed (RTE foods) prior to consumption.138,146 Other sources of NoV contamination are shellfish growing waters (e.g., oysters) and irrigation water (e.g., soft fruits and vegetables). NoVs have also been associated with outbreaks after exposure to contaminated drinking water.50,74,99 The NoVs are very stable in the environment, which greatly contributes to viral spread and disease.40 Studies in human volunteers in the 1970s showed that NV particles remained infectious after the virus had been exposed to pH 2.7 for 3 hours or incubated at 60°C for 30 min.40

6.1.5  Diagnosis Despite numerous attempts, NoV can still not be grown in cell culture32 and although successful passage of NoV in a 3-D cell culture system has been reported,128 this work has not been confirmed by others.139 After the discovery of NV by EM, this method was used for the next 20 years by many research laboratories for the detection of SRSV or NoVs in human fecal samples. Performing EM, however, requires expensive equipment and is a relative insensitive method as it requires at least 106 particles/ml).40 Further developments allowed the visualization of the NoVs by using immune transmission EM63,65,66 or viral antigens by ELISA. Although these refinements in diagnostic procedures increased the sensitivity, broadly reactive antibodies are not widely available.69,123 The cloning and sequencing of several complete NoV genomes (Norwalk and Southampton) followed by numerous reports on partial sequences from other SRSV strains allowed for the development of the first generation of broadly reactive primers to detect NoVs by RT-PCR.2,59,141 These assays allowed detection of NoVs in human fecal samples and using different primer combinations in a nested-PCR format,39 also detection in food matrices such as shellfish. However, since most food matrices contain RT-PCR inhibitors which are coextracted with viral RNA, a critical step in the detection of low-copy number of NoV in foods is the removal of these inhibitors using multi-step elution, concentration, and extraction procedures.6 Over the last 10 years, significant progress

77

has been made in the development of protocols for the improved detection of enteric viruses in shellfish.6,12,23,27,77,125 While different protocols have successfully detected NoVs in other food matrices,69,80 rigorous evaluation of these protocols is needed before standardized protocols can be implemented in laboratories performing routine detection of NoVs in foods. After the initial sampling, most methods for the detection of viruses in food include the following three steps: (1) elution and concentration of the virus particles from the foodstuff, (2) nucleic acid extraction and, (3) detection and confirmation of viral RNA (4). Various strategies for each step have been described15,30,35,86,117,119 and an overview of these methods and their performance characteristics is summarized in Table 6.1. 6.1.5.1  Issues Concerning Sample Processing Sample collection. When salads, soft fruits or whole shellfish are analyzed, generally 10–50 g of sample is analyzed. Since shellfish accumulate viruses in their digestive tissue,103,115 and NV has been shown to bind specifically to carbohydrate structures on this tissue, a number of methods analyzing only this part of shellfish have been published. This approach allows processing of only 1.5–2 g of digestive tissue which has been demonstrated to be sufficient for the recovery and detection of NoV RNA43,77,85 and requires less processing time.6 However, while the dissection of digestive tissue has several advantages for larger shellfish species like oysters and large mussels, this method is technically difficult for small species, such as clams or cockles. As the number of virus particles in each shellfish will often vary, it is important that the sample to be analyzed will represent a number of different shellfish individuals. Elution and concentration of NoV from foods. Elution and concentration of virus particles from the food matrix are the most critical steps of sample processing before RT-PCR detection of viral RNA. Although efforts are aimed to develop horizontal methods applicable to a wide variety of food matrices such as bivalve shellfish, salads, and soft fruits, some specific adjustments of the method may be required to enhance virus elution from a given foodstuff. Therefore, the elution step of virus from food matrices depends on the type of food being analyzed, as well as its state (e.g., fresh, frozen, etc.). In addition, the selection of an elution step needs to be based upon the differences in surface morphology and hydrophobic interactions, internal tissue composition (i.e., RT-PCR inhibitory substances, pH, etc.), whether the virus is present inside (e.g., shellfish flesh) or at the surface of the food (e.g., salads and soft fruit and RTE foods) and conditions of processing to which the food matrix has been subjected previously.1,21,26,45,71 Some methods for shellfish have used acidic adsorption prior to virus elution, but most methods have performed virus elution using a large volume of buffer (two to five times the sample weight). Glycine buffers containing proteins such as beef extract at an alkaline pH (range 8.5–10) have been most commonly used for this purpose. Moreover, when analyzing acidic and pectin containing raspberries or strawberries, the

20 (0.3 g)

8 (0.12 g)

1.5c

1.5c 1.5c 1.5c 1.5c 2c 10c 5c

GuSCN Alkaline

6

4 (0.32 g)

10

50

8

Lettuce

1 (2.5 g) 5 (1.25 g) 5 (0.07 g)

Alkaline Gycine Chlorof-but, CatFloc

Glycine, chlorof Chlorof-but, CatFloc

50 25 1.5c

Glycine, cat-Floc Glycine Glycine Glycine Threonineb Glycine Glycine

Gycine-threonine Chlorof-but, CatFloc Chlorof-but, CatFloc Zirconia beads Proteinase K Trizol ProteinaseK

Clams

20 (1.5 g) 5 (0.1 g)

1 (1 g) 8 (0.8 g)

0.2 (0.04 g)

Viral Elution

Chlorof-but, CatFloc

Glycine Waterb Glycine Sonication Glycine b

20 50 100 10 25 25c 2c

6 (0.1 g) 6 (0.09 g) 0.5 (0.01 g) 0.8 (0.08 g) 1 (0.05 g)

2 (0.4 g)

2 (0.5 g) 1 (0.5 g) 5 (1 g)

Final % Analyzed

25 50 18 50 25

Sample Size (g)

Mussels

Oysters

Food Sample(s)

PEG

Absorbtion to   hydroxyapatite PEG

Ultracentrifugation PEG

Ag capture PEG Ultracentrifugation PEG PEG Ultracentrifugation Ultracentrifugation

PEG PEG Ultracentrifugation

PEG

PEG PEG Ultracentrifugation PEG PEG

Concentration

Table 6.1 Selected Methods Used for Virus Detection in Shellfish, Vegetables and Berries

Trizol QIAshedder™ homogeneiser RNEasy kit

CTAB

Poly A Nucleospin RNAkit RNEasy kit

QIAamp Guanidium, CsCl GuSCN RNEasy kit GuSCN Tryzol + Boom GuSCN

GuSCN+ QIAamp Prot. K, CTAB QIAamp kit RNEasy kit GuSCN GuSCN High Pure Viral NA kit

Prot. K, CTAB

QIAamp Boiling GuSCN GuSCN Tri-reagent

RNA Extraction

50 RT-PCRU NoV 10 PFU HAV 10 RT-PCRU NoV

1.2 TCID50/g HAV 600 PCRU/g HAV

102–103 RT-PCRU NoV

<102 genome copies

22 RT-PCR50U NoV 27 PFU HAV 100 RT-PCRU NoV 100 PFU HAV

1.2 PFU/g EV

Detection Limit(s) (Per Quantity of Analyzed Product)

82

119

37

36 129 22

84 24 110 18 105 107 49

9 122 108 91 60 12 43

6

125 19 106 39 68

Reference

78 Molecular Detection of Foodborne Pathogens

25

Vegetables

15

Raspberries Strawberries Blueberries

20 (3 g)

4 (2.4–3.6 g) Tris-glycine. beef extract

Alkaline Sodium bicarbonate+Soya protein Ultra-filtration

Ultracentrifugation

PEG

Ultracentrifugation

PEG

PEG

Immunomagnetic separation, positively charged filters filtration

Immunomagnetic separation

Magnetic capture

PEG

Filtration

PEG

QIAampa (or miniMAGb)

QIAamp

RNEasy Kit

QIAamp

RNEasy kit

Prot. K, Phenolchlorof CTAB

TrizolTM

QIAamp

Trizol-chlorofpoly(dT) magneticbeads

54 RT-PCRU NoV 12 RT-PCRU HAV 26a or 0b RT-PCRU RV

103–104 RT-PCRU or 102-103 TCID50 HAV

102 RT-PCR U NoV

10% recovery

102 RT-PCRU NoV

10 PFU HAV

0.5 PFU HAV

10-2-102 PFU or 10-105 RT-PCRU HAV

13

118

7

117

7

80

10

123

109

31 30

1.2 × 103 PCRU NoV 50 TCID50 HAV

44

50 PFU HAV

1 RT-PCRU NoV 1 TCID50 HAV

Source: Modified from Le Guyader, F.S., and Atmar, R.L., In Human Viruses in Water, Volume 17: Perspectives in Medical Virology, 1st ed., Bosch, A. (Ed.), Elsevier Science. Amsterdam, 2007. a Detection limit in infectious viruses was calculated taking into account the infectious titer of the HAV stock given by ATCC. b Elution performed after acidic adsorption. c Only digestive tissues analyzed Abbreviations: Glycine, glycine buffer; Chlorof, chloroform; but, butanol; PEG, polyethylene glycol precipitation; ultracentri, ultracentrifugation; prot. K, proteinase K; GuSCN, guanidinium thiocyanate; EV: enterovirus, NoV: norovirus, HAV: hepatitis A virus.

60–90

Alkaline Tris glycine beef-extract

50

Raspberries Strawberries

Alkaline

Gycine Chlorof-but, CatFloc

60

2 (0.2 g)

PBS

PBS

Alkaline glycine

Alkaline

PBS

Alkaline Tryptose phosphate-glycine

Alkaline Tris-glycine-beef extract

10

Raspberries

10

0.4 (0.1 g)

10 (2.5 g)

16 (4 g)

50

One piece of food

25

Green onion Strawberry rinses

Lettuce Strawberries

25

Lettuce Green onions Raspberries

100

25

Green onions

Noroviruses 79

80

release of viruses may be favored by the addition of Tris to the elution buffer, hence increasing the buffer capacity followed by pH neutralization and incubation with pectinase in order to prevent jelly formation.13,30,117,118 After elution of viruses from the food matrix, clarification of the water-phase by extracting with a chloroform-butanol mixture is a rapid step before concentrating the viruses using polyethylene glycol (PEG) precipitation,30 ultracentrifugation,117 or ultrafiltration.13 However, analyzing the digestive tissues of shellfish by direct lysis of the virus particles by proteinase K, or Trizol of the shellfish tissues or by beating with Zirconia beads in the presence of denaturing buffer are elution strategies that have been used successfully.43,60,85,92 A disadvantage of this direct lysis approach is that it often limits the quantity of shellfish tissues that can be analyzed and therefore reduces the chances to recover low amounts of the viral RNA. On the other hand, since small amounts of viruses can still be successfully recovered in relatively smaller amount of digested shellfish, it may be due to the simultaneously decrease in amount of inhibitor which are known to interfere with virus recovery in many concentration methods. There is often an inverse relationship between the recovery of NoV from shellfish tissue and the increasing complexity of the recovery method.43 RNA extraction. Based on the type of food, a variety of methods to extract viral RNA while concurrently reducing the level of PCR inhibitors have been reported4,43 and they are all dependant on the characteristics of the priory concentrated virus suspension. Lysis of the concentrated viruses pellet by proteinase K, followed by purification of nucleic acid using phenol-chloroform and cethyl trimethyl ammonium bromide (CTAB) precipitation was one of the first successful methods described for detection of NoV in shellfish.5 Although this method is quite labor-intensive the efficiency to detect viruses in shellfish has been confirmed in several studies.6,77,78 However, most methods today are based on guanidium extraction either using a variation of the method described by Boom et al.11 or using commercially available kits, based on similar chemistry (QIAamp or RNeasy kit by Qiagen®, or NucliSens magnetic kit by bio-Mérieux). All these RNA extraction methods have proven to be successful for NoV detection in naturally contaminated shellfish, berries, and other food matrices.13,60,80,81,90,114,116 6.1.5.2 Issues Concerning Amplification, Detection, and Quality Assurance (QA) Amplification and detection. The first generation of NoV primers104,144 were not broadly reactive due to the limited knowledge of the tremendous genetic diversity among NoVs. With the availability of more NoV sequences, broadlyreactive RT-PCR assays were designed based on a highly conserved region (region A) within the RdRp.3,12,28,34,76,77,107,141 Although sequences obtained from these assays can be used to distinguish NoV strains, far more reliable genotyping is achieved by amplifying partial regions of the 5′-end of ORF2 (region C) or 3′-end (region D).70,140 While most of the above mentioned RT-PCR assays utilize a virus-specific primer in the RT reaction, random hexamers have also been used successfully to generate cDNA.17,20,39,60 Although nested PCR

Molecular Detection of Foodborne Pathogens

or boosted PCR protocols can increase the sensitivity up to 10–1,000 times12,28,34,39,75,107 extreme care needs to be taken for performing such assays on a routine basis because of the high risk of cross-contamination. Although most RT-PCR assays to detect NoV RNA have been successfully used, no assay which detects all strains has been reported. As a consequence multiplex approaches involving several primer sets and probes are required to address the diversity of these viruses, and no single assay stands out as the best by all criteria.4,143 For example, several outbreak investigations report the use of different primer sets targeting different areas of the genome to be able to amplify the strain in both clinical or environmental samples.79,81,124 Confirmation of RT-PCR amplicons. Although still used in some laboratories, detection of RT-PCR products in ethidium bromide stained agarose gels (i.e., a band having a similar size to that of the positive control) is not considered as an appropriate method for the confirmation of ­conventional RT-PCR assay results.4 Either probe hybridization assays including dot blot hybridization, liquid hybridization, and Southern blot hybridization have been used for confirmation but the limitation is the genetic variability of NoVs which makes it difficult to select a panel of probes to detect most if not all possible NoV strains.76,78 To resolve this and also to attempt to genotype strains, reverse line blot (RLB) assays have been developed for the confirmation of NoV amplicons derived from the polymerase region142 and capsid region.102 These methods are very useful in large-scale epidemiological studies for the initial NoV confirmation of amplicons. To date, most laboratories use TaqMan realtime RT-PCR assay for the detection and confirmation of NoV (see below) and sequencing of region C or region D amplicons is used to determine the genotype or to link geographical different clusters of NoV illness to the same source (e.g., batch of contaminated oysters). Based on analyzing the complete genomic sequence of 18 NoV strains,67 the ORF1–ORF2 junction region has been identified as the most conserved region of the NoV genome. Consequently, GI, and GII specific real-time (Q)RT-PCR assays targeting this region have been developed.61,130,136 These assays have been refined and have been shown to detect NoV in both clinical and environmental samples robustly.25,60,93,108 A selection of primers and probes and different combinations in which they have been used are shown in Table 6.2. Compared to conventional RT-PCR assays, Q-RT-PCR assays is more sensitive, faster, and quantitative if RT-PCR inhibitors are not present in the sample.93 Q-RT-PCR assays also avoid post RT-PCR confirmation steps, such as gel electrophoresis and hybridization or sequencing because positive products can be confirmed by melting curve analysis (in SYBR green Q-RT-PCR) or by probes (in Taqman Q-RT-PCR). In one study, broadly reactive NoV GI and GII TaqMan RT-PCR assays were developed and evaluated on a panel of 84 archived stool samples representing 19 different NoV genotypes and on cDNA from 38 naturally contaminated shellfish samples.60 The sensitivity of these assays is <10 copies of viral genome per reaction. Replacing the degenerated

81

Noroviruses

Table 6.2 Selected Real-Time RT-PCR Primers and Probes Targeting the ORF1-ORF2 Junction for the Detection of Noroviruses in Foods NoV Genogroup NoV GI

NoV GII

NoV IV

a b c

d e

Primers and Probea

Sequence (5’–3’)b

Positionc

Reference

QNIF4 (+) NV1LCR (–) NVGG1p (+)

CGCTGGATGCGNTTCCAT ccttagacgccatcatcatttac FAM-TGGACAGGAGAYCGCRATCT-TAMRAd

5291–5308 5354–5376 5321–5340

25 130

COG1F (+) COG1R (–) Ring 1(a)-TP (–) Ring 1(b)-TP (–)

CGYTGGATGCGNTTYCATGA CTTAGACGCCATCATCATTYAC FAM-AGATYGCGATCYCCTGTCCA-TAMRAd FAM-AGATCGCGGTCTCCTGTCCA-TAMRAd

5291–5310 5354–5375 5321–5340 5321–5340

61

JJV1F (+) JJV1R (–) JJV1P (+)

GCC ATG TTC CGI TGG ATG TCC TTA GAC GCC ATC ATC AT FAM-TGT GGA CAG GAG ATC GCA ATC TC-BHQ

5282–5299 5377–5358 5319–5341

60

Cog 1F (+) Cog 1R (–) Ring 1A (–) Ring 1B (-)

cgYtggatgcglttYcatga cttagacgccatcatcattYac FAM-agatYgcgatcYcctgtcca-BHQe FAM-agatcgcggtctcctgtcca-BHQe

5291–5310 5354–5375 5321–5340 5321–5340

136

QNIF2 (+) COG2R (–) QNIFs (+)

ATGTTCAGRTGGATGAGRTTCTCWGA TCGACGCCATCTTCATTCACA FAM-AGCACGTGGGAGGGCGATCG-TAMRAd

5012–5037 5080–5100 5042–5061

93 61

COG2F (+) COG2R (–) RING2-TP (+)

CARGARBCNATGTTYAGRTGGATGAG TCGACGCCATCTTCATTCACA FAM-TGGGAGGGCGATCGCAATCT-TAMRAd

5003–5028 5080–5100 5048–5067

61

NV2LCF (+) NV2LCR (–) NV2LCpr (+)

GARYCIATGTTYAGRTGGATG TCGACGCCATCTTCATTCAC FAM-TGGGAGGGSGATCGCRATCT-TAMRAd

5006–5026 5081–5100 5048–5067

130

COG2Fex (+) COG2R (–) RING2(a)+RING2 (b) (+)

MRSTGGATGMGRTTYTCWGA TCGACGCCATCTTCATTCACA FAM-TGGGAGGGYGATCGCAATCT (15 pmol)+ TGGGAGGGGGATCGCGATCT (5 pmol)-TAMRAd

5018–5037 5080–5100 5048–5067

62

COG2F (+) NV117 (–) TM3probe (+)

CAR GAR BCN ATG TTY AGR TGG ATG AG TCG ACG CCA TCT TCA TTC AC FAM -TGG GAG GGC GAT CGC AAT CTG GC-TAMRAd

5003–5028 5081–5100 5048–5070

61 51

JJV2F (+) COG2R (-) RING2-TP (+)

CAAGAGTCAATGTTTAGGTGGATGAG TCGACGCCATCTTCATTCACA FAM-AGCACGTGGGAGGGCGATCG-TAMRAd

5003–5028 5080–5100 5048–5067

60

Mon 4F (+) Mon 4R (–) Ring 4 (+)

tttgagtcYatgtacaagtggatgc tcgacgccatcttcattcaca FAM-tgggagggggatcgcgatct-BHQe

718–742 795–815 763–783

136

+, virus sense; −, antivirus sense. Mixed bases in degenerate primers and probes are as follows: Y = C or T; R = A or G; M = A or C; W = A or T; S = C or G; B = not A; I = inosine; N = any. Nucleotide position are based on the Norwalk (GI); accession number M87661, Camberwell virus (GII); accession number AF145896 and the Fort Lauderdale virus (GIV); accession number AF414426. TAMRA, 6-carboxytetramethylrhodamine quencher dye; FAM, 6-carboxyfluorescein reporter dye. BHQ, black hole quencher dye; FAM, 6-carboxyfluorescein reporter dye.

forward primer61 with a nondegenerated primer made this assay significantly more sensitive and at least as sensitive as the nested PCR.60 In another study, a one-step real-time RT-PCR assay was developed for GII and evaluated with the previously described

GI assay61 on an archived panel of 150 naturally contaminated shellfish samples.93 While the GII assay allowed detection of a large range of different strains circulating in France since 1995, the GI strains were detected infrequently. The detection limit as determined on artificially contaminated oysters

82

were found to be 70 and seven RT-PCR detectable units of GI and GII reference NoV strains, respectively.93 Quality assurance (QA)/quality control (QC). QA/QC measures include the use of positive and negative controls to identify false negative or false positive results, respectively. Most false negative results are a consequence of inefficient virus and/or nucleic acid extraction or the presence of inhibitors. Most false positives results are due to crosscontamination. Process extraction controls. Although numerous methods for the detection of viruses in shellfish have been published, it is difficult to compare assay performance between methods. Reasons for this include the use of different virus strains, different foods or species of shellfish, different RT-PCR methods, and the absence of reports of direct comparisons of different methods. The different methods described in Table 6.1 accomplish inhibitor removal to varying degrees, although no systematic evaluation of the efficiency of inhibitor removal has been performed. In order to facilitate the comparison of methods, a universal standard is needed. For this purpose, Costafreda et al.22 proposed the use of Mengo virus as an extraction control to evaluate virus concentration and nucleic acid extraction efficiency. With the accumulating knowledge of the recent cultivable Murine NoV showing similar environmental and chemical resistance as human NoV14 this could be a well suited and perhaps better candidate to be used as an extraction control for NoV. RT-PCR internal controls (IC). Since the concentration of RT-PCR inhibitors varies depending on the type of samples (e.g., species of shellfish, season of harvesting),125 the addition of an internal control like a viral RNA sequence in NoV RT-PCR reactions is critical to realize false negative results as well as providing data on the extraction efficiency of the assay.6,121 Known quantities of RNA transcripts have been spiked into nucleic acid extracts to allow quantitative22,43 or qualitative81,121 assessment of amplification efficiency. Dilution of the viral RNA is another approach to minimize and assess the potential problem of inhibitors.93 However, this approach also affects the assay sensitivity since a smaller amount of shellfish tissue is analyzed. Amplification inhibition seems to be caused by an imbalance between the amount of inhibitors, the amount of virus present in the sample, and the efficiency of the RNA extraction method coupled with its ability to remove inhibitors. Different IC have been used to generate a PCR product with a different size than the target product making it possible to distinguish between the two during gel electrophoresis. However limitations are often the lack of versatility due to amplification by primers that are not broadly reactive. An internal standard was created by cloning a 319 nt sequence of the polymerase gene containing a 156 nt cDNA insert, resulting in a 475 nt RT-PCR product that was amplified by the broadly reactive primer-pair 289/290 that can detect both NoV and sapovirus strains.33 In another study, synthetic short RNA fragments corresponding to the real-time RT-PCR sequence of the primers and probes targeting the ORF1-ORF junction were created based on NoV GI and GII

Molecular Detection of Foodborne Pathogens

reference strains. In each RNA sequence, a restriction site (BamHI) was included to discriminate the sequence from sample RNA if contamination was suspected.25 When using a competitive IC RNA, there are three possible outcomes of RT-PCR assays: (i) no product (the assay has failed, inhibition may be present); (ii) the IC RNA, but not the virus fragment is obtained (the assay has worked, and the sample is most likely negative or viral RNA is below the detection limit); (iii) the virus fragment, with or without the IC RNA fragment, is obtained (the assay has worked, and the sample is positive). EU legislation requires the inclusion of virus controls in standardized methods for the detection of viruses in foods. To harmonize this aspect, efforts are made within the CEN/ TC275/WG6/TAG4 (virus detection in foods) to develop standardized viral RNA extraction and detection protocols, However it becomes essential to gain more information about the practical application of this method to product safety assessment within the EU.

6.2  Methods 6.2.1 Reagents and Equipment Reagents and equipments for the sample preparation and RNA extraction are listed in Table 6.3. Reagents for real-time RT-PCR are listed in Table 6.4 and reactions can be carried out in a MX3000 Real-time PCR thermal cycler (Stratagene, France), or equivalent equipment.

6.2.2 Sample Preparation Based on epidemiological outbreak investigation, food items that have most often been associated with NoV outbreaks include bivalve molluscs especially oysters, mussels, and clams, salad crops such as lettuce and green onions and soft fruits such as raspberries, strawberries, and blackberries. All these food have been implicated in foodborne outbreaks and should be considered as primary targets for virological analysis. A positive result of such analysis may prevent the outbreak to continue by helping the risk management during outbreak investigations to take the proper action which may be immobilization or withdrawal of food stocks, or prohibition of harvest. However, RTE foods served in restaurants or institutions and subsequently implicated in outbreaks due to improper care during handling are less meaningful to be analyzed for viruses as they are often not well preserved and most often the causative agent cannot be identified. A comprehensive epidemiological investigation may be able to find the risk food and possible actions may include a recall of a certain batch of foods. Although bottled mineral water never has been identified as the source of NoV infection, contamination of mineral water with pathogenic bacteria has been reported.137,147 As mentioned in the introduction of this chapter, virus detection in food includes the following steps: sampling, viral elution, concentration, viral RNA amplification, and

83

Noroviruses

Table 6.3 Reagents and Equipments Used for Sample Preparation RNA Extraction of Shellfish and Soft Fruits Matrix Soft fruit

Reagent, Equipment and Consumables

Composition and Concentration of Reagent/Supplier

Elution buffer 1

Glycine, Tris, 1% [wt/vol] beef extract pH 9.5±0.2.

HCL Pectinase Elution buffer 2

Pectinex/Sigma. Glycine, Tris, 1% [wt/vol] beef extract pH 7.0±0.2.

Shellfish

Proteinase K solution

3 U/ml; add 20 mg proteinase K (30 U/mg) to 200 ml molecular grade water and shake to mix/Finnzymes, cat no. F-202S.

Soft fruits and shellfish

Lysis buffer Magnetic extraction reagents Magnetic silica

NucliSens/Bio-Mérieux, cat no. 200292, 280134. NucliSens/Bio-Mérieux, cat no. 200293. NucliSens/Bio-Mérieux, cat no. 280133; magnetic silica is included in the magnetic extraction reagents (cat no. 200293). However, additional silica will be needed due to the increased amount (250 µl per sample) used in this method.

Soft fruit

Falcon tube containing a nylon cell strainer of 40-µm pore size

BD Falcon.

Centricon Plus-70 centrifugal filter device

100K NMWL; Millipore.

Shellfish

Sterile shucking knife and heavy duty safety glove Scissors and forceps Pipettes and pipette tips Sterile petri dishes and sterile razor blades Centrifuge tubes (15 ml)

For the opening of shellfish. For the dissection of shellfish.

Soft fruits and shellfish

Centrifuge NucliSens miniMAG instrument Thermoshaker Waterbath Aspirator or equivalent apparatus Shaking incubator or equivalent Vortex mixer (or pulse vortexer) Micro-tubes (1.5 ml) Eppendorf tubes (1.5 ml) or equivalent

For the chopping of digestive tissue.

Bio-Mérieux, cat no. 200305. Operating at 60°C, 1,400 rpm. Operating at 65°C. For removal of supernatant Operating at 37°C. Bio-Mérieux, cat no. 200294.

Source: Butot, S., Putallaz, T., and Sanchez, G., Appl. Environ. Microbiol., 73, 186, 2007; Greening, G.E. and Hewitt, J., Food Anal. Methods, 1, 109, 2008; and Jothikumar, N. et al., Appl. Environ. Microbiol., 71, 1870, 2005.

detection. Since there is no standard horizontal method to detect NoV in food matrices, we describe here a method for shellfish and a method for soft fruits. For shellfish, sample preparation is based on analyzing the digestive glands which after carefully chopping into very small pieces are subsequently lysed in a proteinase K solution to digest the tissue and release the viral RNA from the virus particles.60 This procedure has been used successfully to detect NoV in many shellfish samples epidemiologically linked to outbreaks and can also be used to quantitatively determine the virus concentration in oysters within a working day.43 For soft fruits, the sample preparation is based on elution of viruses from the fruits by alkaline buffer followed by virus concentration by ultra-filtration.13 The procedure can be applied for different types of vegetables, herbs, and fresh or frozen berries. Evaluation of two commercially available kits (Viral RNA minikit and NucliSens magnetic kit) for viral RNA extraction from the soft fruit concentrate showed

similar recoveries.11 Since the NucliSens magnetic kit has also been demonstrated to work well for RNA extraction on the proteinase K suspension of shellfish, this kit is used here for both matrices. For detection of NoV RNA, two monoplex RT-PCR assays using specific primers and probes targeting the ORF1–ORF2 junction region have been used successfully.25,93 Both assays have been evaluated on a comprehensive panel of stool samples and have been applied effectively to detect NoV GI or GII in naturally contaminated water or shellfish samples.25,93 The overall sample processing scheme is outlined in Figure 6.1. For each sample preparation set-up, it is crucial to include a suitable internal extraction control in order to assure that no cross-contamination has occurred and to monitor the efficiency of the extraction. As a negative control water can be used, while a positive control should be an aliquot of an RNA virus with known concentration and which can be detected by real-time RT-PCR.

84

Molecular Detection of Foodborne Pathogens

Table 6.4 TaqMan Real-Time Protocol for Detection of NoV GI and GII in Shellfish and Soft Fruits Reagent UltraSense reaction mix* UltraSense enzymes mix* ROX* RNase inhibitor QNIF4 GI forward primer NV1LCR GI reverse primer NV1LCpr GI probe QNIF2d GII forward primer COG2R GII reverse primer QNIFS GII probe PCR qualified water Total volume/reaction before RNA addition

Concentration/Reaction

Concentration Stock Solution

Volume/Reaction (µl)

1× – 1× – 500 nM 900 nM 250 nM 500 nM 900 nM 250 nM – –

5× – 50× – 50 µM 50 µM 50 µM 50 µM 50 µM 50 µM – –

5.00 1.25 0.50 2U 0.25 0.45 0.13 0.25 0.45 0.13 12.42 20

Source: da Silva, A.K. et al., Appl. Environ. Microbiol., 73, 7891, 2007; and Loisy, F. et al., J. Virol. Methods, 123, 1, 2005. * RNA UltraSense™ One-Step Quantitative RT-PCR System. Invitrogen, cat. no. 11732927.

15 g of soft fruit or vegetables

1.5 g of chopped digestive gland representing at least six shellfish

Lyse with 2 ml proteinase K at 37˚C for 60 min with shaking

Elute by agitating for 15 min in 60 ml elution buffer (50 mM Glycine, 100 mM Tris, 1% (w/v) beef extract pH 9.5) Filtrate through cell stainer Pectinase treatment at neutral pH (for pectin containing berries only)

Centrifugation at 10,000 × g for 10 min

Centrifugation at 3500 × g for 15 min

Ultra-filtration of supernatant at 3500 × g for 15–45 min

Lyse viral capsid and remaining tissue by transferring the entire virus suspension to 4 ml lysis buffer.

Viral RNA extraction using magnetic silica NoV RNA detection by TaqMan real-time RT-PCR

Figure 6.1  Flow chart of norovirus detection in soft fruits and shelfish. (From Butot, S., Putallaz, T., and Sanchez, G., Appl. Environ. Microbiol., 73, 186, 2007; da Silva, A.K. et al., Appl. Environ. Microbiol., 73, 7891, 2007; Jothikumar, N. et al., Appl. Environ. Microbiol., 71, 1870, 2005; Loisy, F. et al., J. Virol. Methods, 123, 1, 2005; Svraka, S. et al., J. Clin. Microbiol., 45, 1389, 2007.)

Soft fruits.13 Weight 15 g of soft fruits into a stomacher bag and add 60 ml of elution buffer 1 (Table 6.3). Shake gently for 15 min at room temperature (Figure 6.1). Transfer the eluate to a Falcon tube containing a nylon cell strainer (Table 6.3) to remove particulate debris. Rinse the berries and the cell strainer with 6 ml and 2 ml of elution buffer 1,

respectively. Adjust the pH of the transferred eluate to pH 7.0 with 9.5 M HCl and add 300 µl of pectinase. Incubate while gently shaking for 30 min at room temperature. Centrifuge the extract at 3,500 × g for 15 min. and transfer the supernatant to a Centricon Plus-70 centrifugal filter device (Table 6.3) and centrifuge at 3,500 × g to concentrate

85

Noroviruses

the virus particles to ca. 400 µl. The centrifugal filter device is subsequently rinsed with 200 µl of elution buffer 2 (Table 6.3 and Figure 6.1). Shellfish. 43,60 Select a suitable number (preferably a minimum of six) of live oysters or mussels. Wash off any mud from the shell and open the shellfish with a clean shucking knife. Ensure the hand holding the shellfish is protected with a heavy duty safety glove. Dissect out the digestive glands using scissors and forceps (or equivalent tools). Transfer the glands to a clean petri dish and chop finely with a razor blade. During preparation the shellfish must be placed on ice. Transfer a 1.5 g portion of chopped glands into a 15 ml centrifuge tube. Add 2 ml of proteinase K solution (Table 6.3) and mix well. Incubate at 37°C in a shaking incubator (or equivalent) for 60 min. Transfer the tube to a water bath at 65°C for 15 min as a second incubation. Centrifuge at 10,000 × g for 10 min, decant the supernatant, measure, and record the volume and retain the sample (supernatant) for nucleic acid extraction. RNA extraction. Add the entire sample obtained from the shellfish (proteinase K suspension) or soft fruits (ultrafiltrate) sample preparation to a tube containing 4.5 ml of NucliSens lysis buffer and vortex briefly. Continue the procedure according to the manufactures instructions except for using 250 µl of well-mixed magnetic silica solution. Resuspend the extracted RNA in 100 µl elution buffer and use this for real-time RT-PCR.

6.2.3 Detection of Noroviruses by Real-time RT-PCR Amplification and inhibition controls. A negative amplification control (water) and positive amplification controls (e.g., GI and GII reference RNA in low copy numbers) should be included in each amplification series in order to monitor for cross contamination, and amplification efficiencies. In addition the inhibitor removal during the nucleic acid extraction and potential amplification inhibition should be assessed by e.g., testing each sample in duplicates of undiluted and 1/10 diluted RNA. Mastermix and RT-PCR conditions. The mastermixes for real-time RT-PCR should be prepared fresh immediately prior to use. A separate mastermix should be prepared for each of the NoV genogroups. Add the ingredients in the order listed in Table 6.4 and prepare enough mastermix for at least one reaction more than required. Dispense 20 µl mastermix to each well plus 5µl of the RNA sample to be tested (giving a total volume of 25 µl in each well). RT should be carried out for 30 min at 50°C, and denaturation for 5 min at 95°C, followed by 45 cycles of PCR amplification (denaturation at 95°C for 15 s, annealing, and extension at 60°C for 1 min).25 Reporting of results. Analyze the amplification plot using the approach recommended by the manufacturer. Given that all the controls are as expected, a sample is considered positive if the curve giving a Ct value has a proper shape. A sample is considered “not positive” if there is no Ct value or

if the curve does not have the proper shape regardless of the Ct value.

6.3  Conclusion and Future Perspectives NoVs are now recognized as the most important foodborne pathogens contributing to more outbreaks than all other known bacterial, viral, and protozoan pathogens combined. Therefore, efforts to prevent and control viral foodborne illnesses should be primarily directed toward NoVs. Foodborne transmission of NoV involves RTE food items contaminated with fecal material or vomit. Another important source of NoV contamination is water used for food washing or irrigation as well as shellfish growing waters contaminated by sewage infiltration or direct fecal contact. Which foods are implicated in a NoV outbreak is typically identified based on an epidemiological outbreak investigation where the relative risk of exposure to different food items is calculated. Standardized methods for the detection of viruses in foods other than shellfish are not yet available because until recently the methods used to recover and detect viruses from these foods were inadequate. However, the recent development of sensitive TaqMan-based real-time RT-PCR methods have greatly improved the sensitivity and specificity of the detection of NoVs but standardized upstream methods such as concentration and extraction methods have not been established. Using a cocktail of HBGAs that have been recognized as potential receptors of NoVs, or specific monoclonal antibodies to pre-enrich the virus particles regardless of the type of food, may lead to robust standardized protocols to detect one of the most important groups of foodborne pathogens.

Acknowledgments This work was supported by the Danish Veterinary and Food Administration (DVFA), the European Union Project BIOTRACER (FOOD-2006-CT-036272) and the National Food Institute, Technical University of Denmark (DTU). The authors would like to thank Soizick Le Guyader, Laboratoire de Microbiologie, IFREMER, Nantes, France for helpful discussions.

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7 Rotaviruses

Dongyou Liu, Larry A. Hanson, and Lesya M. Pinchuk Mississippi State University

Contents 7.1 Introduction......................................................................................................................................................................... 91 7.1.1 Classification and Biology....................................................................................................................................... 91 7.1.1.1 Rotaviruses................................................................................................................................................ 91 7.1.1.2 Structural and NSPs.................................................................................................................................. 91 7.1.1.3 Serogroups, Serotypes, and Genotypes..................................................................................................... 92 7.1.1.4 Biology...................................................................................................................................................... 92 7.1.2 Clinical Manifestation, Pathogenesis, and Epidemiology....................................................................................... 93 7.1.2.1 Clinical Manifestation............................................................................................................................... 93 7.1.2.2 Pathogenesis.............................................................................................................................................. 93 7.1.2.3 Epidemiology............................................................................................................................................ 94 7.1.3 Diagnosis.................................................................................................................................................................. 94 7.1.3.1 Phenotypic Techniques.............................................................................................................................. 94 7.1.3.2 Molecular Techniques............................................................................................................................... 95 7.2 Methods............................................................................................................................................................................... 95 7.2.1 Sample Preparation ................................................................................................................................................. 95 7.2.2 Detection Procedures............................................................................................................................................... 96 7.2.2.1 RT-PCR Detection of Rotavirus G and P Genotypes............................................................................... 96 7.2.2.2 Real-time RT-PCR Detection of Rotaviruses Groups A and C................................................................ 96 7.3 Conclusion........................................................................................................................................................................... 97 References.................................................................................................................................................................................... 98

7.1  Introduction 7.1.1 Classification and Biology 7.1.1.1 Rotaviruses The genus Rotavirus encompasses a group of nonenveloped, segmented, double-stranded (ds) RNA viruses in the family Reoviridae. Being spherical particles measuring about 72 nm in diameter with a smooth outer edge, the rotaviruses are so named because their icosahedral shell or capsid (with 32 capsomers) forms a rim round the nucleic acid core, giving a wheel-like appearance.1 The rotavirion capsid is composed of three concentric protein layers encasing a genome of 11 unique dsRNA segments (amounting to 16–21 kb in total), with each segment usually coding for a single structural or nonstructural protein (NSP) (with the exception of segments 9 and 11, which each code for two proteins). The inner (core) capsid protein layer encasing dsRNA comprises structural protein VP2 and small numbers of structural proteins VP1 and VP3. The middle capsid protein layer consists of structural protein VP6, which defines rotavirus serogroup (A–G) specificity and is the most immunogenic protein in rotavirus infection. The outer capsid layer is made up of structural protein VP7 (a glycoprotein) in which the protease-sensitive structural protein VP4 spikes are embedded. Accounting

for rotavirus serotype (neutralization)–specificity, the VP4 protein (encoded by segment 4) is P serotype antigen, while the VP7 protein (encoded by segment 7) is G serotype antigen.2,3 7.1.1.2 Structural and NSPs In total, six structural proteins (i.e., VP1, VP2, VP3, VP4, VP6, and VP7) and six NSPs (i.e., NSP1, NSP2, NSP3, NSP4, NSP5, and NSP6) are encoded by rotaviruses. Specifically, rotavirus VP1, VP2, VP3, VP4, VP6, and VP7 are encoded by their corresponding gene segments, NSP1, NSP2, NSP3, NSP4, NSP5, and NSP6 are encoded by segments 8, 7, 10, 11, and 11, respectively. In other words, NSP5, and NSP6 are encoded in overlapping reading frames of a single segment 11.2,4 Located in the inner capsid layer linked to the dsRNA core, VP1, VP2, and VP3 along with other NSPs play a key role in viral transcription. Specifically, being an RNA polymerase enzyme, VP1 is responsible for producing mRNA transcripts for the synthesis of viral proteins and for replicating the rotavirus genome RNA segments for new virus particles inside the infected host cells. VP2 binds the RNA genome; and VP3 is a guanylyl transferase that catalyses the formation of the 5′ cap in the post-transcriptional modification of mRNA. This 91

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process stabilizes viral mRNA by protecting it from ­nucleases of the host cell. Situated in the middle capsid layer, VP6 forms the bulk of the capsid with a role in transcription. Being common to all rotaviruses, and highly antigenic, VP6 has been targeted for classification and serological identification of rotaviruses, helping separate them into serogroups A, B, C, D, E, F, and G. Protruding as a spike on the surface of the virion, VP4 binds to molecules on the surface of host receptor cells, and thus facilitates the viral entry into the cell. Prior to the virus turning infectious, VP4 needs to undergo modification (into VP5 and VP8) with the help of a protease enzyme found in the gut. Neutralizing antibodies to VP4 can be used to determine the P-type (P for protease-sensitive) and this protein helps determine the virulence of the virus. The outer layer of the virion is formed by VP7. This is a glycoprotein that is targeted by some neutralizing antibodies and is used to provide the antigenic G-type classification (G for glycoprotein) of the strain. Together with VP4, VP7 generates neutralizing, and protective antibodies against subsequent infections.3 NSP1 (encoded by segment 5), NSP2 (encoded by segment 8), and NSP3 (encoded by segment 7) are RNA-binding proteins that are required either for genome replication, shutdown of cellular protein synthesis, or translational regulation. NSP4 is a viral enterotoxin that increases the intracellular Ca2+ concentration and disturbs the cellular homeostasis of the host, a main factor contributing to malabsorption and diarrhea. NSP5 is (encoded by segment 11) accumulates in the viroplasm, and NSP6 (also encoded by segment 11) is a nucleic acid binding protein. 7.1.1.3 Serogroups, Serotypes, and Genotypes Rotaviruses in the genus Rotavirus can be divided into seven serogroups (A–G) in accordance with serological reactions between the structural protein VP6 (located in the middle capsid layer) and corresponding antisera. Further, rotaviruses are differentiated into 16 G-genotypes and at least 27 P genotypes based on assessment of the outer capsid layer structure proteins VP7 (which defines G-serotypes/genotypes as it is glycoprotein) and VP4 (which defines P-serotypes/genotypes as it is a protease sensitive protein) as well as their underlying genes.3 To designate a P-genotype, a number in square brackets is added after “P” (e.g., P[8]); and to indicate a P-serotype, a serotype number (without bracket) is attached to “P” followed by the genotype in brackets (e.g., P2A[6]). Because there are greater numbers of P-type genotypes than serotypes, P-type antigens are denoted with their genotype referenced in brackets (e.g., P[4], P[8]). G-serotypes are numbered similarly, and the G-genotype number is the same as the G-serotype. Thus, rotavirus G-serotypes are often referred to by their serotypes alone (e.g., G1, G2, G3, etc). Given that the VP4 and VP7 proteins are encoded by separate gene segments, new rotavirus P–G serotype antigen combinations may be generated through reassortment after dual (or mixed) infection of single cells. In human

Molecular Detection of Foodborne Pathogens

infections, rotavirus strains of G–P combinations G1P[8], G2P[4], G3P[8], G4P[8], G9P[6], and G9P[8] are routinely detected, although those bearing rare or unusual G- and/or P-genotypes (e.g., G5, G8, G10, G11, G12, P[11], P[14], and P[25]) have also been described.5 Due to the ubiquitous presence of rotaviruses in the animal kingdom, interspecies transmission and genetic reassortment events make the emergence of novel rotavirus strains from existing animal and human strains a common event.6 In another scheme of classification, the 11 segments of rotaviruses are separated by polyacrylamide gel electrophoresis (PAGE), and a characteristic visual pattern known as RNA electropherotype is generated for individual serogoups.7–9 For example, serogroup A rotaviruses displays a 4–2–3–2 segment migration pattern (indicating the number of rotavirus gene segments of similar sizes that migrate at similar speed on acrylamide gels); serogroup B rotaviruses show a 4–2–2–3 pattern and serogroup C rotaviruses have a 4–3–2–2 pattern.8 This information is valuable since detection of different electropherotypes during an outbreak may indicate the possible emergence of novel or recurring virus strains. Further, on the basis of the migration rate of segment 11 on acrylamide gels in relation to segment 10, long or short electropherotype of rotaviruses can be distinguished. That is, while the standardsized segment 11 of long-electropherotype strains moves faster than segment 10, the short-electropherotype phenotype with a partial duplication in segment 11 causes it to migrate more slowly than gene segment 10.4,10,11 Sequencing is being used more frequently in recent years with VP4, VP6, and VP7 initially used for genotyping. Later the enterotoxin gene NSP4 was added and most recent classification scheme is the use of sequence data from all 11 gene segments.9 Although all rotavirus serogroups (A–G) are infective to animals, only serogroups A–C are found in human infections. Indeed, the majority of human rotavirus isolates share a common group antigen A, which is further divided into at least seven serotypes, and is the single most important cause of severe acute gastroenteritis among infants and young children worldwide. Serogroups B–G rotaviruses are morphologically identical but serologically unrelated to serogroup A. Rotavirus B causes diarrhea in adults (thus called adult diarrhea rotavirus or ADRV) as well as in individuals of other age groups Rotavirus C is responsible for sporadic cases of diarrhea in children. Group D, F, and G mainly infect poultry and Group E may infect swine.8 7.1.1.4 Biology Rotaviruses are tolerant of a range of humidity and pH (3–9), and are unaffected by ether, chloroform, quaternary ammonium disinfectants, and sodium hypochlorite.8 However, they show a reduced infectivity at higher temperatures and are vulnerable to ethanol, phenol, formalin, 37% formaldehyde (1:10), 0.75% hexachlorophene (1:3), and 67% chloramine-T (1:5).8 Thus, rotaviruses are resistant to sanitary measures which are adequate for eliminating bacteria and parastes, and they are stable in the environment. It is no surprise

Rotaviruses

therefore that rotaviruses are able to remain infectious under the inclement and hostile conditions associated with mammalian gastrointestinal tract prior to entry into the epithelial cells of small intestinal villi and subsequent replication in the cytoplasm of these cells. Rotaviruses are capable of undergoing constant genetic changes via sequential point mutations, genetic reassortment, genomic rearrangement, or intragenic recombination.5,12–26 As rotaviruses possess 11 dsRNA segments encoding distinct structural and NSPs, different strains of human and/ or animal origins often exchange genetic materials through reassortment and evolve into new viral species, serotypes, and genotypes.27–33 This is demonstrated by the observations that the common genotype P[8] strains with serotype G1, G3, G4, or G9 VP7 genes have the same genome constellation (Wa genogroup), and except VP7, all other genes are able , to cross-hybridize with other genotype members. The generation of new P–G genotype combinations by the introduction of genes from novel serotypes represents another useful way to generate rotavirus diversity. Another major source of human rotavirus diversity involves the introduction of animal rotavirus genes either through transmission of whole viruses or through reassortment.34 Coinfections with two different or more rotavirus serotypes may also contribute to a great degree of strain diversity among rotaviruses.

7.1.2 Clinical Manifestation, Pathogenesis, and Epidemiology 7.1.2.1  Clinical Manifestation Rotavirus enters the host via fecal-oral route, and upon survival in the acidic environment of the host’s gastrointestinal tract, it infects the epithelial cells of small intestine villi and secretes an enterotoxin, which is responsible for gastroenteritis with diarrhea and sometimes dehydration-related death as a consequence.35 The clinical symptoms of rotavirus gastroenteritis often appear two days after infection, starting with vomiting followed by four to eight days of profuse, watery diarrhea accompanied with low grade fever, headache, malaise, nausea, or cramping. Although rotavirus gastroenteritis is considered as a self-limiting disease, without appropriate intervention, a combination of watery diarrhea, vomiting, and fever often can lead to rapid dehydration and ultimately death. While rotavirus infection in newborn children is often associated with mild or asymptomatic disease, it causes the most severe symptoms in children six months to two years of age, the elderly, and those with suppressed immune functions. Most adults are not susceptible to rotavirus infection due to immunity acquired in childhood, and gastroenteritis in adults may be attributable to nonrotavirus infection. However symptomatic reinfections may occur with a different rotavirus A serotype. Upon resolution of gastroenteritis symptoms, rotavirus antigen may be shed in the stools for approximately one week but for many months in immunocompromised individuals. In fact, the number of infectious rotavirus particles in the stool of an infected person can be higher than

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10 trillion per gram; and only ten to 100 of these particles are sufficient to transmit infection to another person. 7.1.2.2  Pathogenesis Principally transmitted by the fecal-oral route via contact with contaminated hands, surfaces, objects, food, and water and possibly by the respiratory route, rotavirus relies on its triple capsid layers of proteins to avoid elimination by the acidic pH of the stomach and the proteolytic enzymes in the gut. At the same time, cleavage, and removal of its outer capsid layer by intestinal proteases such as chymotrypsin facilitates its efficient colonization and enhances its infectivity.3 The virus first enters the epithelial cells of the villi in the duodenum by receptormediated endocytosis and forms an endosome. Sialic acid or galactose-based receptors as well as coreceptors like integrins may be involved in endocytosis.3 In addition, the virus may also gain entry into the absorptive enterocytes by direct penetration.3 Then, through the coordinated action of the outer capsid layer proteins (VP7 and the VP4 spike), the virus disrupts the membrane of the endosome, resulting in intracellular calcium imbalance, breakdown of VP7 trimers into single protein subunits and formation of a double-layered particle (DLP) by the remaining capsid layer proteins VP2 and VP6 covering viral dsRNA in the core. In doing so, the viral RNA in the core avoid triggering innate host immune responses called RNA interference. Once outside the endosome, the virus multiplies in the epithelial cells of the small intestinal villi and causes structural and functional changes.1 The viral RNA-dependent RNA polymerase generates mRNA transcripts of the double stranded viral genome, and assembles into DLPs in viroplasm (made by NSP5 and NSP2) around the cell nucleus where most of the rotavirus proteins accumulate. After migrating to the endoplasmic reticulum, DLPs acquire their outer capsid layer (formed by VP7 and VP4) and become infectious viruses which are released from the cell upon lysis.37 As one of the two main types of enterocytes lining the small intestine (the other being crypt cells) villus enterocytes are mature, nonproliferating cells covering the villi with either digestive or absorptive functions. The absorptive enterocytes express disaccharidases, peptidases, and other enzymes on the apical surface to carry out their digestive functions. This is achieved by both passive diffusion of solutes along electrochemical or osmotic gradients and active transport, which employs sodium-glucose cotransporter 1 (SGLT1) to transport water along with solute in addition to passive transportation along osmotic gradients. The crypt cells covering the crypts are the progenitors of the villus enterocytes. Without the well-defined microvilli and absorptive functions, crypt cells secrete Cl− ions into the intestinal lumen. Thus, the absorptive villi and secretive crypts maintain a constant bidirectional flux of electrolytes and water across the epithelium. When rotavirus begins to replicate in the nondividing mature enterocytes near the tips of the villi, it secretes a viral enterotoxin, NSP4, which alters calcium-dependent cell permeability, elevates the chloride secretion and causes cell lysis.1,3,37 The viral NSP4 is then released to extracellular area

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where it binds to adjacent cells. Thus, together with enterocyte ­destruction, rotavirus infection decreases absorption of Na+, water, and mucosal disaccharidases, and this malabsorption causes undigested mono- and disaccharides, carbohydrates, fats, and proteins to remain in the colon, which are discharged with unabsorbed water. Thus, a decrease in the absorptive functions of villus enterocytes, an increase in the secretive functions of crypt enterocytes and subsequent destruction of villus entorocytes contribute to malabsorption, leading to diarrhea. In the later stages of rotavirus infection, enterocytes in the villi regenerate and the patient regain normal body weight within ten to 28 days after infection. Thus, rotavirus infection is considered a self-limiting disease as long as the dehydration is not severe as to causing death of young ones.8 It has been shown that the CD8+ cytotoxic T lymphocytes (CTL) as well as CD4+ helper T cells (TH) are instrumental in helping clear the rotavirus infection.37,38 7.1.2.3 Epidemiology Group A rotavirus is a ubiquitous pathogen that accounts for >90% of rotavirus gastroenteritis in humans, resulting in millions of cases of diarrhea, almost 2 million in hospitalization and an estimated 611,000 in death.39–63 It is the leading single cause of severe diarrhea among infants and children (from six months to five years old), with boys being twice as likely to be admitted to hospital as girls. However, group A rotavirus is not considered to cause any significant degree of disease in adults. By contrast nongroup A rotaviruses (i.e., B and C) are much less common and only cause sporadic family and community outbreaks of gastroenteritis in all age groups.64 Primary infections with group A rotavirus peak between nine and 23 months of age, with infants of six to 23 months of age having the greatest risk for being hospitalized. By five years of age, most children have exposure to the virus with or without symptoms, as evidenced by the presence of specific serum antibodies against the pathogen.65 Although adults with exposure to the virus are frequently asymptomatic, they may still shed the viral particles in their stools, which can be transmitted to children. The elderly and individuals with compromised immune functions are at significant risk of developing severe diarrhea and showing extended period of viral dissemination, for whom medical intervention is often required. Other factors that may affect the severity of rotavirus infections in humans include host variations, geographic/seasonal differences, and emergence of new virus strains, etc. For example, malnutrition in humans often increases the severity of rotavirus diarrhea, delays small intestinal recovery, and modifies intestinal inflammatory responses. Rotavirus infections occur primarily in the winter in temperate areas, but they occur throughout the year in the tropics; due possibly to the seasonal changes in temperature and humidity. The close contact between humans and domestic animals promotes the emergence of novel virus strains and varieties, as rotaviruses have the capacity to exchange genetic materials via reassortment, leading to novel viral identities that are more potent and are unrecognized by host immune network. Additionally, the increased consumption of ready-to-eat or raw food products

Molecular Detection of Foodborne Pathogens

(e.g., seafood and vegetables) also facilitates transmission of animal rotaviruses to humans, as group A rotaviruses are shed in large numbers in stools (>1012 particles/g feces), which may pollute waters used for farming and bivalve shellfish culture.66,67

7.1.3  Diagnosis 7.1.3.1  Phenotypic Techniques Rotaviruses are capable of growing in cell culture in primary calf kidney cells as well as in established cell lines such as MA104, MDBK or PK-15, which facilitate their identification and characterization. Other phenotypic procedures include electron microscopy, PAGE-based electropherotyping, enzyme linked immunosorbent assay (ELISA), dot-immunoblotting, immuno-fluorescence test, immunoperoxidase test, passive hemagglutination assay (HA), latex agglutination test, and dot blot hybridization, etc.68 Rotavirus serogroups A–G are determined by serological assays with antibodies raised against structural protein VP6 located in the middle capsid layer; and rotavirus group A serotypes are similarly assessed by using antibodies to structural proteins from outer capsid layers (i.e., a glycoprotein VP7 defining G serotypes/genotypes and a protease sensitive protein VP4 defining P serotypes/genotypes). As a consequence, four common serotypes (designated as serotypes G1–G4), corresponding to antigenic determinants located on the VP7 protein, have been identified in rotavirus group A. These four serotypes (G1–G4) represent at least 90% of rotavirus group A strains circulating globally. Other 12 human rotavirus G serotypes (e.g., G8, G9, and G12) are considered rare. Given that about 30% of rotavirus-positive stool specimens have nontypeable (NT) strains, which cannot be typed with existing monoclonal antibodies, use of culture and other techniques (e.g., molecular analysis) is essential to characterize these NT strains. Laboratory diagnosis of human rotavirus infections is possible by enzyme immunoassay for detection of antibodies against rotavirus VP6 group A-specific antigen in diarrheal stools. Another approach is to employ anti-VP6specific antibodies immobilized onto latex (reverse passive latex agglutination test), nitrocellulose, or nylon (rapid chromatographic assays) for the immunological capture of rotavirus VP6 antigen in feces, and subsequent detection with a second labeled, types-specific anti-VP6 antibody, and chromatographic assays, or by visible agglutination of latex particles. On the whole, rotavirus-specific immunoglobulin (Ig) A or IgG antibodies are more readily detected than rotavirus antigen in fecal specimens. Thus, the antigen detection techniques often result in an underestimation of rotavirus disease and asymptomatic infection. Nonetheless, immunological assays such as ELISA are dependent on the presence of significant amounts of intact rotavirus proteins in stool samples for consistent detection. Due to the frequent digestion or degradation of these proteins by endogenous and exogenous proteolytic enzymes, together with the possibility of the rotavirus

Rotaviruses

strains bearing point mutation particles, application of typespecific monoclonal antibodies in immunological assays for rotaviruses in stool may not always give reliable account on the true status of human rotavirus gastroenteritis.69 7.1.3.2  Molecular Techniques Molecular procedures including reverse-transcription PCR (RT-PCR), nested/multiplex RT-PCR, restriction fragment length polymorphism (RFLP) and real-time reverse­transcription PCR provide a much specific, sensitive, and rapid approach for detection and genotyping of rotaviruses.70–77 Other molecular methods such as nucleotide sequencing and oligonucleotide microarray hybridization offer the opportunity to discriminate mixed rotavirus infections.78–80 PCR assays have been applied since the early 1990s for the detection and identification of human and animal rotavirus VP4 and VP7.81,82 Apart from showing the predominance of only four common rotavirus G serotypes (G1–G4) among children, these techniques facilitated the identification of at least 43 distinct P–G type combinations among the ten human rotavirus G serotypes and 11 rotavirus HRV P serotypes and subtypes. Use of a multiplexed, seminested RT-PCR method permitted the detection of both the common G serotypes (G1–G4) and the rare serotypes (e.g., G5, G8, and G9), and the genotyping of common P types (P[4] and P[8]) and rare P types (P[6], P[9], and P[10]). Given that large numbers of rotaviruses are shed in stools (>1012 particles/g feces), and discharged in sewage, it is no surprise that rotavirus is one of the common viruses present in waters and shellfish. Although food- or waterborne outbreaks of rotavirus gastroenteritis are infrequent, the presence of rotaviruses in water, vegetables, and seafood imply a health risk to humans.83 Due to the fact that a relative low density of rotavirus is found in water and shellfish specimens, various sample processing procedures are needed to concentrate the virus and to remove of natural inhibitors to DNA polymerase.83 Direct alkaline elution84 and acid adsorption– neutral elution85 are examples of the common and efficient methods used to concentrate viruses from whole shellfish. Using the acid adsorption–alkaline elution and reconcentration using speedVac centrifugation together with an expanded rotavirus microarray assay of Chizhikov et al.78 Honma et al.86 identified from (i) five clinically relevant human rotavirus VP4 genotypes (P[4], P[6], P[8], P[9], and P[14]) and (ii) five additional human rotavirus VP7 genotypes (G5, G6, G8, G10, and G12) on one chip. Use of a microtiter plate hybridization-based ELISA coupled with PCR (PCRELISA) also facilitated the identification of clinically relevant G (G1–G6, G8–G10, G12) and P (P[4], P[6], P[8], P[9], and P[14]) genotypes of group A rotavirus.

7.2  Methods 7.2.1 Sample Preparation Oysters. Oyster samples often contain low levels of rotaviruses, which need to be concentrated before RT-PCR

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detection. A useful technique to prepare oyster samples is the acid adsorption-alkaline elution method.83,85 Briefly, oyster samples are transported to the laboratory in chilled containers. On arrival, the oysters are washed, scrubbed, and the shells opened with a sterile shucking knife. The liquor or mantle fluid is drained into a discard container. The oyster meat, except the adductor muscle that was left attached to the shell, is collected, cut into small pieces, and trimmed to 25 g for analysis of rotavirus. Seven volumes of chilled, sterilized distilled water are added to 25 g of oyster flesh and homogenized at high speed twice using a blender (Hamilton Beach, Southern Pines, NC). The conductivity of the homogenate is reduced to less than 2000 μS/cm by adding sterilized distilled water. The homogenate of oyster is adjusted to pH 4.8 with 1 N HCl, shaken for 15 min, and centrifuged at 2000 × g for 20 min at 4°C. The supernatant is discarded and the pellet is suspended in 25 ml of 2.9% tryptose phosphate broth (TPB) containing 6% glycine, pH 9.0 for elution of the virus, shaken for 15 min, and centrifuged at 10,000 × g for 15 min at 4°C. The supernatant (S1) is collected and the pellet is reeluted with 25 ml of 0.5 M arginine–0.15 M NaCl, pH 7.5. The suspension is shaken for 15 min and centrifuged. The supernatant (S2) is collected, combined with S1 and adjusted to pH 7.2 with 1 N HCl. The virus in the supernatant is precipitated by adding 12.5% PEG 8000 and 0.3 M NaCl. The mixture is refrigerated overnight at 4°C. The pellet is dissolved in 15 ml of 0.05 M phosphate-buffered saline (PBS), pH 7.5 and precipitated again with PEG–NaCl. The mixture is stirred for 2 h at 4°C and then centrifuged at 10,000 × g for 10 min. The pellet is dissolved in 5 ml of PBS. The virus is extracted with chloroform at a final concentration of 30%. After centrifugation at 3000 × g for 10 min, the top layer of the aqueous phase is collected (S3). The pellet at the interface between the solvent and the aqueous phase is re-extracted with 0.5 volume of arginine–NaCl, pH 7.5. After centrifugation, the top layer (S4) is collected and combined with S3. The sample is reconcentrated using speedVac centrifugation to reduce the volume of the concentrate to approximate 1  ml and stored at −80°C until nucleic acid extraction. Viral RNA from oyster samples is then extracted using RNeasy® mini kit (Qiagen AG, Basel, Switzerland). In brief, 200 μl of the concentrated oysters is lysed and RNA is purified on the silica-based column according to the manufacturer’s protocol. RNA bound to the membrane in the column is eluted in 60 μl of warm RNase-free water.87 Another method for preparation of oyster samples is by direct alkaline elution.84 Briefly, seven volumes of chilled, sterilized, distilled water are added to the oyster flesh and homogenized using a blender (Hamilton Beach, Southern Pines, NC). The virus is eluted from oyster tissues with 10% TPB containing 0.05 M glycine, pH 9.0 and re-eluted with 0.5 M arginine–0.15 M sodium chloride (NaCl), pH 7.5. The pooled eluates are precipitated twice by adding polyethylene glycol (PEG 8000) and NaCl to a final concentration of 12.5% PEG–0.3 M NaCl. After dissolving the pellet, the virus is extracted with chloroform to a final concentration of 30% and re-extracted with 0.5 volume of arginine–NaCl, pH 7.5.

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Molecular Detection of Foodborne Pathogens

The aqueous phase is collected and kept at −80°C prior to RNA extraction. Stool. The viral ds RNA is extracted from stools or infected cell culture lysates using the TRIzol method (Invitrogen, Carlsbad, CA). Alternatively, total nucleic acids can be extracted from stool samples using a single protocol.74 Briefly, fecal specimens are prepared as 10% suspensions with STAR buffer (stool transport and recovery buffer; Roche Diagnostics). Chloroform aliquots of 0.1× volume are added, and following mixing, the samples are centrifuged at 8000 × g for 10 min. The aqueous layer (stool extract) is removed to a  fresh tube and stored at −80°C, or the extraction is completed before freezing. Stool extracts of 200 μl are further purified with the Qiagen QIAamp DNA Total, and nucleic acids are eluted from the spin columns using 50 μl of nuclease-free water. Extracted nucleic acids are stored at −80°C.74

7.2.2  Detection Procedures 7.2.2.1 RT-PCR Detection of Rotavirus G and P Genotypes Santos et al.88 described a RT-PCR for detection of group A rotavirus G and P genotypes. The viral dsRNA is extracted from stools or infected cell culture lysates using the TRIzol method and subjected to reverse transcription followed by RT-PCR. The cDNA of the rotavirus VP7 or VP4 gene is synthesized by RT-PCR using primers that are biotin labeled at their 5′ ends. Four different VP7 gene-specific primer pairs are selected to increase the efficiency of VP7 gene amplification (Table 7.1). Initially, the Beg9–End9 primer pair, which amplifies the entire VP7 segment of most of rotavirus strains, is used. When the VP7 gene is not amplified with this primer pair, one of the following three primer pairs is used: sBeg9– RVG9, 9Con2–G922, or LID1–G922. The sBeg9–RVG9 primer pair is used as the first choice because it provides the best amplification. For VP4 gene amplification, two different primer pairs are selected (Table 7.2). The first primer pair used is Con3–Con2. For those strains for which the VP4 gene is negative with this primer pair, the primer pair F4–C8 is used for amplification (Table 7.2).

Table 7.2 VP4 Primers for RT-PCR Detection of Rotavirus P Genotypes Primer

Sequence

Positions

Con3

5′-TGGCTTCGCTCATTTATAGACA-3′

Con2

5′-ATTTCGGACCATTTATAACC-3′

11–32

F4

5′-TATAAAATGGCTTCIYTVAT-3′

4–23

C8

5′-ATTGGIGTYTGATAITCRTC-3′

1446–1467

868–887

Procedure: (1) Prepare RT-PCR mixture (50 µl) containing 1 × reaction mix (a buffer containing 0.2 mM of each deoxynucleoside triphosphate (dNTP), 2 mM MgSO4), SuperScript™ III RT/Platinum® Taq Mix (Invitrogen, Life Technologies, Carlsbad, CA), 0.25 µM each primer pair, distilled water to 48 µl and 2 µl extracted RNA. (2) Subject the genomic RNAs to one cycle of reverse transcription (25°C at 5 min followed by 42°C at 45 min). (3) Perform PCR amplification for 30 cycles of 94°C for 30 sec, 40°C for 1 min and 72°C for 1.5 min; and a cycle of 72°C for 5 min. (4) Analyze the PCR products by agarose gel electrophoresis and visualize by staining with ethidium bromide. Note: application of the VP7 and VP4 primers in RT-PCR permits detection of rotavirus G and P genotypes. To further distinguish among rotavirus G–P combinations, specific probes can be then utilized in a PCR-ELISA system.88 7.2.2.2 Real-time RT-PCR Detection of Rotaviruses Groups A and C Logan et al.89 described a real-time RT-PCR for detection of rotavirus A, and rotavirus C from stool specimens, with primers and probes targeting rotavirus group A structural protein VP6 gene and rotavirus group C structural protein VP7 gene (Table 7.3). Procedure:

Table 7.1 VP7 Primers for RT-PCR Detection of Rotavirus G Genotypes Primer Sequence Positions Beg9 1–28 5′-GGCTTTAAAAGAGAGAATTTCCGTCTGG-3′ End9 1036–1062 5′-GGTCACATCATACAATTCTAATCTAAG-3′ sBeg9

5′-GGCTTTAAAAGAGAGAATTTC-3′

1–21

RVG9

1044–1062

5′-GGTCACATCATACAATTCTAATCTAAG-3′

LID1

5′-ATGTATGGTATTGAATATACCA-3′

48–70

9Con1

5′-TAGCTCCTTTTAATGTATGG-3′

37–56

G922

5′-GTRTARAAIACTTGCCACCA-3′

922–941

(1) Prepare reverse transcription mixture (40 μl) containing 1 × RT buffer, 5.5 mM MgCl2, 0.5 mM each dNTP, 2.5 μM random hexamer, 0.4 U/μl RNase inhibitor, 1.25 U/μl Multiscribe reverse transcriptase (TaqMan reverse transcription kit, Applied Biosystems), and 17.5 μl of extracted nucleic acids. (2) Immediately before testing, remove extracted nucleic acids (prepared by Qiagen QIAamp DNA kit, see above) from −80°C storage and add to 200μl thin-walled PCR tubes containing the RT buffer, dNTPs, and MgCl2.

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Rotaviruses

Table 7.3 Primers and Probes for Real-Time RT-PCR Detection of Rotavirus Groups A and C Target gene

Primer or probe

Concentration (μM)

Position

Rotavirus A (VP6)

RotaA-fwd1

5′-GGATGTCCTGTACTCCTTGTCAAAA-3′

0.60

26–50

RotaA-fwd2

5′-GGAGGTTCTGTACTCATTGTCAAAAA-3′

0.60

26–51

RotaA.rev1

5′-TCCAGTTTGGAACTCATTTCCA-3′

0.60

170–149

RotaA.rev2

5′-TCCAGTTTGAAAGTCATTTCCATT-3′

0.60

170–147

5′-(VIC)-ATAATGTGCCTTCGACAAT-3′

0.25

93–75 93–72

RotaA.probe1 RotaA.probe2 Rotavirus C (VP7)

Sequence

5′-(VIC)-AATATAATGTACCTTCAACAAT-3′

0.25

RotaC.fwd

5′-TTAGATACTACAAGTAATGGAATCGGATGT-3′

0.60

655–684

RotaC.rev

5′-TGGGTGTCATTTGATACAACTTCA-3′

0.60

731–708

RotaC.probe

5′-(FAM)-CAGCTAGTACAGAAACTT-3′

0.20

689–706

(3) Denature the reaction components by heating at 95°C for 5 min on a GeneAmp 9700 thermocycler (Applied Biosystems) and then snap-chill on ice for 5 min. Add random hexamer, RNase inhibitor, and Multiscribe reverse transcriptase to the reaction tube. (4) Conduct reverse transcription on a GeneAmp 9700 thermocycler as follows: one cycle of 25°C for 10 min; one cycle of 42°C for 30 min; one cycle of 48°C for 20 min; and one cycle of 95°C for 5 min. Store the completed RT products at −20°C until use. (5) Prepare real-time PCR mixture (25-μl) containing 1 × TaqMan Universal Mastermix (containing AmpErase uracil-N-glycosylase, with dTTP partially replaced by dUTP), the appropriate forward and reverse primer(s) and MGBNFQ probe(s) at concentrations as detailed in Table 7.3 and 2 μl of template RT products. (6) Perform real-time PCR on an ABI 7000 sequence detector (Applied Biosystems) using universal thermal cycling conditions: one cycle of 50°C for 2 min (dUTP glycosylase step); one cycle of 95°C for 10 min; and 40 cycles of 95°C for 15 sec, 60°C for 1 min. Use sequence detection software version 1.2 for all data analysis. Note: rotaviruses serogroups A, B, and C are known to cause gastrointestinal infections in humans. Group A rotaviruses are predominant and result in severe diarrheal diseases in infants and young children. Group B rotaviruses cause adult diarrhea and are reportedly geographically confined, having been first identified in a large waterborne epidemic in China. Group C rotaviruses, an emerging cause of gastroenteritis in children over two years old and in adults, have now been identified as causative agents of gastroenteritis in both sporadic cases and outbreaks worldwide. Use of this real-time RT-PCR permits simultaneous detection of rotaviruses serogroups A and C in stool specimens.89

7.3  Conclusion Rotaviruses are dsRNA viruses that are separated into seven groups (A–G) based on serological reactions bet­ ween the structural protein VP6 (located in the middle capsid layer) and corresponding antisera. While serogroups A–G are infective to animals, only serogroups A, B, and C are known to cause gastrointestinal infections in humans. Group A rotaviruses are predominant and result in severe diarrheal diseases in infants and young children. Group A rotaviruses can be classified into at least 27 P genotypes on the basis of their VP4 protein and 16 G genotypes on the basis of their VP7 protein. Group B rotaviruses cause adult diarrhea and are reportedly geographically confined. Group C rotaviruses, an emerging cause of gastroenteritis in children over two years old and in adults, have now been identified as causative agents of gastroenteritis in both sporadic cases and outbreaks worldwide. Due to their distinct genetic structures (with 11 gene segments) and their ability to undergo genetic changes via sequential point mutations, genetic reassortment, genomic rearrangement, or intragenic recombination, new rotaviruses may emerge constantly. This has necessitated the development and application of rapid and accurate laboratory assays for detection and typing of rotaviruses. While immunological assays such as ELISA have proven valuable for the diagnosis of rotavirus infections, these phenotypic procedures suffer from poor sensitivity and inability to detect rotaviruses secreting mutated target proteins. Application of molecular procedures such as PCR and microarray has made rapid, sensitive, and precise detection and typing of rotaviruses feasible. Indeed, these new generation techniques facilitate the characterization of rotavirus strains not commonly found in humans and animals. Considering the existence of large number of genotypes within group A rotavirus, use of standard RT-PCR is cumbersome as multiple individual reaction tubes may have to be set up for determining genotypes that are very close to other similar genotypes. By combining RT-PCR with microarray or ELISA, it is possible to type all rotavirus G and P genotypes in an efficient manner. Further adoption of these RT-PCR

98

assays into real-time platforms will enhance laboratory detection and identification of disease-causing rotaviruses.

References







1. Estes, M.K. and Kapikian, A.Z. Rotaviruses. In: Fields Virology, 5th edn., vol 2. pp. 1917–1974. Knipe, D.M. et al. (eds.). Lippincott Williams & Wilkins, Philadelphia, PA, 2007. 2. Desselberger, U. Rotavirus: basic facts. In: Rotaviruses Methods and Protocols, pp. 1–8. Gray, J. and Desselberger, U. (eds.). Humana Press, 2000. 3. Desselberger, U., Gray, J. and Estes, M.K. Rotaviruses. In: Topley and Wilson’s Microbiology and Microbial Infections, pp. 946–958. Mahy, B.W.J. and Meulen, V.T. (eds.). ASM Press, USA, 2005. 4. Mohan, K.V. and Atreya, C.D. Nucleotide sequence analysis of rotavirus gene 11 from two tissue culture-adapted ATCC strains, RRV and Wa. Virus Genes, 23, 321, 2001. 5. Griffin, D.D. et al., Surveillance of rotavirus strains in the United States: identification of unusual strains. J. Clin. Microbiol., 38, 2784, 2000. 6. Gentsch, J.R. et al. Serotype diversity and reassortment between human and animal rotavirus strains: implication for rotavirus vaccine programs. J. Infect. Dis., 192, S146, 2005. 7. Maunula, L. and von Bonsdorff, C.H. Short sequences define genetic lineages: phylogenetic analysis of group A rotaviruses based on partial sequences of genome segments 4 and 9. J. Gen. Virol., 79, 321, 1998. 8. Steele, A.D., Geyer, A. and Gerdes, G.H. Rotavirus infections. In: Infectious Diseases of Livestock, pp. 1256–1264. Coetzer, J.A.W. and Tustin, R.C. (eds.). Oxford University Press, Southern Africa, 2004. 9. Matthijnssens, M. et al. Recommendations for the classification of group A rotaviruses using all 11 genomic RNA segments. Arch. Virol., 153, 1621, 2008. 10. Kobayashi, N. et al. Unusual human rotavirus strains having subgroup I specificity and “long” RNA electropherotype. Arch. Virol., 109, 11, 1989. 11. Matsui S.M. et al. Sequence analysis of gene 11 equivalents from “short” and “super short” strains of rotavirus. J. Virol., 64, 120, 1990. 12. Adah, M.I. et al. Nigerian rotavirus serotype G8 could not be typed by PCR due to nucleotide mutation at the 3′ end of the primer binding site. Arch. Virol., 142, 1881, 1997. 13. Gouvea, V. and Brantly, M. Is rotavirus a population of reassortants? Trends Microbiol., 3, 159, 1995. 14. Desselberger, U. Genome rearrangements of rotavirus. Adv. Virus Res., 49, 69, 1996. 15. Alfieri, A.A. et al. Characterization of human rotavirus genotype P[8]G5 from Brazil by probe-hybridization and sequence. Arch. Virol., 141, 2353, 1996. 16. Suzuki, Y., Gojobori, T. and Nakagomi, O. Intragenic recombination in rotaviruses. FEBS Lett., 427, 183, 1998. 17. Diwarkarla, C. and Palombo, E. Genetic and antigenic variation of capsid protein VP7 of serotype G1 human rotavirus isolates. J. Gen. Virol., 80, 341, 1999. 18. Unicomb, L.E. et al. Evidence of high-frequency genomic reassortment group A rotavirus strains in Bangladesh: emergence of type G9 in 1995. J. Clin. Microbiol., 37, 1885, 1999. 19. Griffin, D.D. et al. Characterization of nontypeable rotavirus strains from the United States: identification of a new rotavirus reassortant (P2A[6], G12) and rare P3[9] strains related to bovine rotaviruses. Virology, 294, 256, 2002.

Molecular Detection of Foodborne Pathogens 20. Iturriza-Gómara, M. et al. Reassortment in vivo: driving force for diversity of human rotavirus strains isolated in the United Kingdom between 1995 and 1999. J. Virol., 75, 3696, 2001. 21. Iturriza-Gómara, M., Kang, G. and Gray, J. Rotavirus genotyping: keeping up with an evolution population of human rotaviruses. J. Clin. Virol., 31, 259, 2004. 22. Arista, S. et al., G2 rotavirus infections in an infantile population of the South of Italy: variability of viral strains over time. J. Med. Virol., 77, 587, 2005. 23. Awachat, P.S. and Kellar, S.D. Unexpected detection of simian SA-11-human reassortant strains of rotavirus G3P[8] genotype from diarrhea epidemic among tribal children of Western India. J. Med. Virol., 77, 128, 2005. 24. Rahman, M. et al. Typing of human rotaviruses: nucleotide mismatches between the VP7 gene and primer are associated with genotyping failure. Virol. J., 2, 24, 2005. 25. Parra, G.I. and Espinola, E.E. Nucleotide mismatches between the VP7 gene and the primer are associated with genotyping failure of a specific lineage from G1 rotavirus strains. Virol. J., 3, 35, 2006. 26. Chan-it, W. et al. Multiple combinations of P[13]-like genotype with G3, G4, and G5 in porcine rotaviruses. J. Clin. Microbiol., 46, 1169, 2008. 27. Isegawa, Y. et al. Determination of bovine rotavirus G and P serotypes by polymerase chain reaction. Mol. Cell. Probes, 7, 277, 1993. 28. Gouvea, V., Santos, N. and Timenetsky, M.C. VP4 typing of bovine and porcine group A rotaviruses by polymerase chain reaction. J. Clin. Microbiol., 32, 1333, 1994. 29. Gouvea, V. et al., Identification of two lineages (Wa-like and F45-like) within the major rotavirus genotype P[8]. Virus Res., 59, 141, 1999. 30. Gosh, S. et al. Molecular characterization of a porcine group A rotavirus strain with G12 genotype specificity. Arch. Virol., 151, 1329, 2006. 31. Gulati, B.R. et al. Diversity in Indian equine rotaviruses: identification of genotype G10, P6[1] and G1 strains and a new VP7 genotype (G16) strain in diarrheic foals in India. J. Clin. Microbiol., 45, 972, 2007. 32. Martella, V. et al. Identification of a novel VP4 genotype carried by a serotype G5 porcine rotavirus strain. Virology, 346, 301, 2006. 33. Martella, V. et al. Identification of group A porcine rotavirus strains bearing a novel VP4 (P) genotype in Italian swine herds. J. Clin. Microbiol., 45, 577, 2007. 34. Mascarenhas, J.D. et al. Detection of a neonatal human rotavirus strain with VP4 and NSP4 genes of porcine origin. J. Med. Microbiol., 56, 524, 2007. 35. Parashar, U.D. et al. Rotavirus and severe childhood diarrhea. Emerg. Infect. Dis., 12, 304, 2006. 36. Chauhan, R.S., Dhama, K. and Mahendran, M. Pathobiology of rotaviral diarrhea in calves and its diagnosis and control: a review. J. Immunol. Immunopathol., 10, 1, 2008. 37. Ramig, R.F. Pathogenesis of intestinal and systemic rotavirus infection. J. Virol., 78, 10213, 2004. 38. Hoshino, Y. and Kapikian, A.Z. Rotavirus serotypes: classification and importance in rotavirus epidemiology, immunity and vaccine development. J. Health Popul. Nutr., 18, 5, 2000. 39. Guanasena, S. et al. Relative frequency of VP4 gene alleles among human rotaviruses recovered over a 10-year period (1982–1991) from Japanese children with diarrhea. J. Clin. Microbiol., 31, 2195, 1993.

Rotaviruses 40. Das, B. K. et al. Characterization of rotavirus strains from newborns in New Delhi, India. J. Clin. Microbiol., 32, 1820, 1994. 41. Cunliffe, N.A. et al. Rotavirus G and P-types in children with acute diarrhea in Blantyre, Malawi, from 1997 to 1998: predominance of novel P[6] G8 strains. J. Med. Virol., 57, 308, 1999. 42. Cunliffe, N.A. et al. Rotavirus strain diversity in Balantyre, Malawi, from 1997 to 1999. J. Clin. Microbiol., 39, 839, 2001. 43. Iturriza-Gómara, M. et al. Molecular epidemiology of human group A rotavirus infections in the UK between 1995 and 1998. J. Clin. Microbiol., 38, 4094, 2000. 44. Ramachandran, M. et al. Molecular characterization of serotype G9 rotavirus strains from a global collection. Virology, 278, 436, 2000. 45. Banyai, K. et al. Detection of human rotavirus serotype G6 in Hungary. Epidemiol. Infect., 130, 107, 2003. 46. Banyai, K. et al. Eight-year survey of human rotavirus strains demonstrates circulation of unusual G and P types in Hungary. J. Clin. Microbiol., 42, 393, 2004. 47. Kirkwood, C. et al. Genetic and antigenic characterization of rotavirus serotype G9 strains isolated in Australia between 1997 and 2001. J. Clin. Microbiol., 41, 3649, 2003. 48. Laird, A.R. et al. Characterization of serotype G9 rotavirus strains isolated in the United States and India from 1993 to 2001. J. Clin. Microbiol., 41, 3100, 2003. 49. Martella, V. et al. Detection of emerging rotavirus G9 serotype at high frequency in Italy. J. Clin. Microbiol., 41, 3960, 2003. 50. Martella, V. et al. Nucleotide variation in the VP7 gene affects PCR genotyping of G9 rotaviruses identified in Italy. J. Med. Virol., 72, 143, 2004. 51. Page, N.A. and Steele, A.D. Antigenic and genetic characterization of serotype G2 human rotavirus strains from South Africa from 1984 to 1998. J. Med. Virol., 72, 320, 2004. 52. Nielsen, N.M. et al. Characterization of rotavirus strains among hospitalized and non-hospitalized children in GuineaBissau, 2002. A high frequency of mixed infections with serotype G8. J. Clin. Virol., 34, 13, 2005. 53. Khamrin, P. et al. Emergence of human G9 rotavirus with an exceptionally high frequency in children admitted to hospital with diarrhea in Chiang Mai, Thailand. J. Med. Virol., 78, 273, 2006. 54. Castello, A.A. et al. Molecular epidemiology of group A rotavirus diarrhea among children in Buenos Aires, Argentina, from 1999 to 2003 and emergence of the infrequent genotype G12. J. Clin. Microbiol., 44, 2046, 2006. 55. Pietruchinski, E. et al. Rotavirus diarrhea in children and adults in a southern city of Brazil in 2003: distribution of G/P types and finding of a rare G12 strain. J. Med. Virol., 78, 1241, 2006. 56. Samajdar, S. et al. Changing pattern of human group A rotaviruses: emergence of G12 as an important pathogen among children in eastern India. J. Clin. Virol., 36, 183, 2006. 57. Uchida, R. et al. Molecular epidemiology of rotavirus diarrhea among children and adults in Nepal: detection of G12 strains with P[6] or P[8] and a G11P[25] strain. J. Clin. Microbiol., 44, 3499, 2006. 58. Volotão, E.M. et al. Rotavirus surveillance in the city of Rio De Janeiro-Brazil during 2000–2004: detection of unusual strains with G8P[4] or G10P[9] specificities. J. Med. Virol., 78, 263, 2006.

99 59. Duan, Z.-J. et al. A novel human rotavirus of G5P[6] genotype identified from a Chinese girl with diarrhea. J. Clin. Microbiol., 45, 1614, 2007. 60. Fischer, T.K. et al. Characterization of rotavirus strains in a Danish population: high frequency of mixed infections and diversity within the VP4 gene of P[8] strains. J. Clin. Microbiol., 43, 1099, 2005. 61. Fischer, T.K. et al. Hospitalization and deaths from diarrhea and rotavirus among children <5 years of age in the United States, 1993–2003. J. Infect. Dis., 195, 1117, 2007. 62. Rahman, M. et al. Evolutionary history and global spread of the emerging G12 human rotaviruses. J. Virol., 81, 2382, 2007. 63. Steyer, A.M. et al. Rotavirus genotypes in Slovenia: unexpected detection of G8P[8] and G12P[8] genotypes. J. Med. Virol., 79, 629, 2007. 64. Smith M.J. et al. The clinical and molecular epidemiology of community- and healthcare-acquired rotavirus gastroenteritis. Pediatr. Infect. Dis. J., 27, 54, 2008. 65. Dunn, S.J. et al. Serotypic and genotypic characterization of human serotype 10 rotaviruses from asymptomatic neonates. J. Clin. Microbiol., 31, 165, 1993. 66. Gratacap-Cavallier, B. et al. Detection of human and animal rotavirus sequences in drinking water. Appl. Environ. Microbiol., 66, 2690, 2000. 67. van Zyl, W.B. et al. Molecular epidemiology of group A rotaviruses in water sources and selected raw vegetables in southern Africa. Appl. Environ. Microbiol., 72, 4554, 2006. 68. Malik, S.V.S. et al. Data sheet on rotaviruses (global status of rotavirus infections in man and animals). In: Animal Health and Production Compendium. CAB International, Wallingford, UK, 2005. 69. Coulson, B.S. VP4 and VP7 typing using monoclonal antibodies. Arch. Virol. (Suppl.), 12, 113, 1996. 70. Gouvea, V. et al. Polymerase chain reaction amplification and typing of rotavirus nucleic acid from stool specimens. J. Clin. Microbiol., 28, 276, 1990. 71. Gentsch, J.R. et al. Identification of group A rotavirus gene 4 types by polymerase chain reaction. J. Clin. Microbiol., 30, 1365, 1992. 72. Taniguchi, K. et al. Identification of human and bovine rotavirus serotypes by polymerase chain reaction. Epidemiol. Infect., 109, 303, 1992. 73. Iturriza-Gómara, M. et al. Comparison of specific and random priming in the reverse transcriptase polymerase chain reaction for genotyping group A rotaviruses. J. Virol. Methods, 78, 93, 1999. 74. O’Neill, H.J. et al. Clinical utility of nested multiplex RT-PCR for group F adenovirus, rotavirus and Norwalk-like viruses in acute viral gastroenteritis in children and adults. J. Clin. Virol., 25, 335, 2002. 75. Santos, N. et al. VP7 gene polymorphism of serotype G9 rotavirus strains and its impact on G genotype determination by PCR. Virus Res., 90, 1, 2002. 76. Santos, N. et al. VP7 gene polymorphism of serotype G9 rotavirus strains and its impact on G genotype determination by PCR. Virus Res., 93, 127, 2003. 77. Fischer, T.K., and Gentsch, J.R. Rotavirus typing methods and algorithms. Rev. Med. Virol., 14, 71, 2004. 78. Chizhikov, V. et al. Detection and genotyping of human group A rotaviruses by oligonucleotide microarray hybridization. J. Clin. Microbiol., 40, 2398, 2002. 79. Lovmar, L. et al. Microarray for genotyping human group A rotavirus by multiplex capture and type-specific extension. J. Clin. Microbiol., 41, 5153, 2003.

100 80. Domingues, A.L.S., Silva, M.H. and Gouvea, V. Biotin-psolaren labeled cDNA amplicons for genotyping rotavirus strains by dot hybridization assay. J. Virol. Methods, 140, 228, 2007. 81. Adler, M. et al. Detection of rotavirus from stool samples using a standardized immuno-PCR (“Imperacer”) method with end-point and real-time detection. Biochem. Biophys. Res. Commun., 333, 1289, 2005. 82. Ahmed, K., Nakagomi, T. and Nakagomi, O. Molecular identification of a novel G1 VP7 gene carried by a human rotavirus with a super-short RNA pattern. Virus Genes, 35, 141, 2007. 83. Kittigul, L. et al. An efficient virus concentration method and RT-nested PCR for detection of rotaviruses in environmental water samples. J. Virol. Methods, 124, 117, 2005. 84. De Medici, D. et al. Detecting the presence of infectious hepatitis A virus in molluscs positive to RT-nested-PCR. Lett. Appl. Microbiol., 33, 362, 2001.

Molecular Detection of Foodborne Pathogens 85. Mullendore, J.L., Sobsey, M.D. and Shieh, Y.C., Improved method for the recovery of hepatitis A virus from oysters. J. Virol. Methods, 94, 25, 2001. 86. Honma, S. et al. Development and validation of DNA microarray for genotyping group A rotavirus VP4 (P[4], P[6], P[8], P[9], and P[14]) and VP7 (G1 to G6, G8 to G10, and G12) genes. J. Clin. Microbiol., 45, 2641, 2007. 87. Kittigul, L. et al. Development of a method for concentrating and detecting rotavirus in oysters. Int. J. Food Microbiol., 122, 204, 2008. 88. Santos, N. et al. Development of a microtiter plate hybridization-based PCR-enzyme-linked immunosorbent assay for identification of clinically relevant human group A rotavirus G and P genotypes. J. Clin. Microbiol., 46, 462, 2008. 89. Logan, C., O’Leary, J.J. and O’Sullivan, N. Real-time reverse transcription-PCR for detection of rotavirus and adenovirus as causative agents of acute viral gastroenteritis in children. J. Clin. Microbiol., 44, 3189, 2006.

8 Sapoviruses Grant S. Hansman

National Institute of Infectious Diseases

Contents 8.1 Introduction........................................................................................................................................................................101 8.1.1 Genetic Classification ............................................................................................................................................101 8.1.2 Serological Studies................................................................................................................................................. 103 8.1.3 Cross-Reactive Studies.......................................................................................................................................... 103 8.1.4 Diagnostic Techniques........................................................................................................................................... 103 8.2 Methods............................................................................................................................................................................. 103 8.2.1 Reagents and Equipment........................................................................................................................................ 103 8.2.2 Sample Preparation................................................................................................................................................ 105 8.2.2.1 Primary-treated Domestic Wastewater (Method 1)................................................................................ 105 8.2.2.2 Secondary-treated Effluent (Method 1) and River Water (Method 1)..................................................... 105 8.2.2.3 Influent and Effluent Water (Method 2).................................................................................................. 105 8.2.2.4 River Water (Method 2)........................................................................................................................... 106 8.2.2.5 Seawater.................................................................................................................................................. 106 8.2.2.6 Oysters..................................................................................................................................................... 106 8.2.2.7 Shellfish................................................................................................................................................... 106 8.2.2.8 Stool......................................................................................................................................................... 106 8.2.3 Detection Procedures............................................................................................................................................. 106 8.2.3.1 EM........................................................................................................................................................... 106 8.2.3.2 ELISA Detection..................................................................................................................................... 107 8.2.3.3 RNA Extraction....................................................................................................................................... 107 8.2.3.4 Reverse Transcription.............................................................................................................................. 107 8.2.3.5 Nested PCR............................................................................................................................................. 108 8.2.3.6 Real-time RT-PCR.................................................................................................................................. 108 8.2.3.7 Phylogenetic Analysis............................................................................................................................. 109 8.3 Conclusions and Future Perspectives................................................................................................................................ 109 References.................................................................................................................................................................................. 109

8.1  Introduction Viruses can infect animals, plants, and bacteria. Viruses are composed of either single-stranded or double-stranded nucleic acid, which can be either DNA or RNA. Viral infections in humans usually result in an immune response and disease. In this chapter, we utilize human sapovirus, a cause of gastroenteritis, as a model to illustrate the various ways to prepare viral samples for molecular detection and identification. In recent years, a number of important findings concerning human sapovirus were discovered, which were likely, in part, due to improved detection techniques and increased surveillance. Human sapovirus strains were detected in water samples, which included untreated wastewater specimens, treated wastewater samples, and river samples;1,2 human sapovirus strains were detected in clams destined for human consumption in Japan;3 porcine sapovirus was detected in oysters destined for human consumption in the USA;4 and the number of sapovirus-associated outbreaks of gastroenteritis,

especially involving adults, appears to be steadily increasing, suggesting that sapovirus virulence and/or prevalence may be increasing.5–8

8.1.1 Genetic Classification The virus family Caliciviridae contains four genera ­Sapo­virus, Norovirus, Lagovirus, and Vesivirus, which include Sapporo virus, Norwalk virus, Rabbit hemorrhagic disease virus, and Feline calicivirus, respectively. Human sapovirus is an etiological agent of gastroenteritis. The prototype strain of human sapovirus, the Sapporo virus, was originally discovered from an outbreak in an orphanage in Sapporo, Japan, in 1977.9 In that study, Chiba et al. identified viruses with the typical animal calicivirus morphology, the Star-of-David structure, by electron microscopy (EM). Besides having this classical structure, sapovirus particles are typically 41–46 nm in diameter and have a cup-shaped depression and/or ten spikes on the outline. The sapovirus 101

102

Molecular Detection of Foodborne Pathogens

genome is a ­single-stranded, positive sense RNA molecule of approximately 7.5 kb that is polyadenylated at the 3′ end. Sapovirus can be divided into five genogroups (GI–GV) Figure 8.1), among which GI, GII, GIV, and GV are known to infect humans, whereas sapovirus GIII infects porcine species (Figure 8.1). The sapovirus GI, GIV, and GV genomes are each predicted to contain three main open reading frames (ORFs), whereas sapovirus GII and GIII have two ORFs. Sapovirus ORF1 encodes for nonstructural proteins, including the VPg, protease, and RNA dependent RNA polymerase (RdRp), and a major capsid protein (VP1). Sapovirus ORF2 and ORF3 encoded proteins of yet unknown functions. Recently, naturally occurring intragenogroup recombinant sapovirus strains were identified (i.e., strains Mc10 and C12).10 When polymerase-based grouping was performed, Mc10 and C12 strains clustered together, but when capsid-based grouping

was performed, these two strains belonged in two distinct genotypes. In a more recent study, intergenogroup recombinant sapovirus strains were identified (i.e., strains SW278 and Ehime1107).11 Phylogenetic analysis of the nonstructural region (i.e., genome start to capsid start) grouped SW278 and Ehime1107 strains into GII, while the structural region (i.e., capsid start to genome end) grouped these strains into GIV. By comparing the sequence similarity across the length of the genomes using SimPlot software,12 a potential recombination site was discovered, at a point where the similarity analysis showed a sudden drop in nucleotide identity after the RdRp region, indicating that a recombination event occurred at the RdRp-capsid junction. Since the discovery of these intragenogroup and intergenogroup recombinant sapovirus strains classification numbering schemes have conflicted between the different research groups. Ideally, both polymerase and

1000

1000 943

Mc114 Manchester Sapporo

1000

Dresden

805

Chiba000496 Ehime643 1000

1000

1000

Parkville Houston27 Stockholm Chiba010658

999

Syd3

Arg39 1000 963

Mc2 Bristol SK15

1000

1000 1000 410

GI/2

GI/3

GIV/1

SW278 NK24

1000

469

GI/5

Ehime1107

1000

916

GI/4

Potsdam

1000

918

0.02

Yokote1

1000

536

GI/1

Syd53 C12

GV/1 GII/1 GII/6 GII/3

Mc10

GII/2

Mex340

GII/4

Cruise ship

GII/5

PEC

GIII

Figure 8.1  Phylogenetic tree of sapovirus based upon the entire VP1 nucleotide sequences. Different genogroups and genotypes are indicated. The numbers on each branch indicate the bootstrap values for the genotype. Bootstrap values of 950 or higher were considered statistically significant for the grouping. The scale represents nucleotide substitutions per site. GenBank accession numbers for the reference strains are as follows: Arg39, AY289803; Bristol, HCA249939; C12, AY603425; Chiba000496, AJ412800; Chiba010658, AJ606696; Cruise ship, AY289804; Dresden, AY694184; Ehime643, DQ366345; Ehime1107, DQ058829; Houston27, U95644; Manchester, X86560; Mc2, AY237419; Mc10, AY237420; Mc114, AY237422; Mex340, AF435812; NK24, AY646856; Parkville, U73124; PEC, AF182760; Potsdam, AF294739; Sapporo, U65427; SK15, AY646855; Stockholm, AF194182; SW278, DQ125333; Syd3, DQ104357; Syd53, DQ104360; and Yokote1 AB253740.

103

Sapoviruses

capsid genes should be analyzed. However, this is not always practical, due to amplification difficulties, time and cost of reagents. Therefore, many phylogenetic studies have used the 5′ terminus of the capsid gene.

8.1.2 Serological Studies Most studies agree that sapovirus infection is more frequent in young children than adults and that infection in children almost always occurs by 5 years of age. In addition, children at daycare centers and institutions are at greatest risk of sapovirus-associated infection and transmission. In an early study of sapovirus antibody prevalence in the general community,13 Sakuma et al. indicated that sapovirus infections were acquired more readily after 2 years of age than before 2 years of age, and especially in infants attending nurseries and children attending kindergarten or primary schools. Grohmann et al.14 also found that infants under 6 months of age had the highest rate of sapovirus-associated gastroenteritis. Sapovirus also causes sporadic cases of acute gastroenteritis requiring hospitalization as well as symptomatic and asymptomatic infections not requiring hospitalization in the community.8,14–28 However, only a limited number of sapovirus studies have been conducted, therefore it has been difficult to check for correlations between or to draw conclusions about rates of incidence, detection, and overall prevalence. Epidemiological studies have been conducted in a number of countries, including Australia, Canada, Finland, France, Japan, Mexico, Mongolia, Spain, Sweden, Taiwan, Thailand, UK, US, and Vietnam. The rates of incidence, detection, and overall prevalence of sapovirus infections vary in each country and setting, and are likely affected by the diagnostic techniques used.29

8.1.3 Cross-Reactive Studies Human sapovirus is noncultivable, but expression of a subgenomic-like construct (i.e., VP1 to genome end) or VP1 alone in insect or mammalian cells can result in the formation of virus-like particles (VLPs) that are morphologically and antigenically similar to native sapovirus.30–36 Recently, we compared the antigenic relationships among all human sapovirus genogroups.37–39 Hyperimmune antisera that were raised against the VLPs reacted strongly against the homologous VLPs. However, several antisera weakly cross-reacted against heterologous VLPs using an antibody ELISA. On the other hand, an antigen ELISA showed that the sapovirus VLPs belonging to all human genogroups were antigenically distinct. These findings provide evidence of a likely correspondence between the sapovirus antigenicity and VP1 genogrouping and genotyping.

8.1.4  Diagnostic Techniques Several groups have used enzyme immunoassays (EIAs) to screen for sapovirus antibodies.20,25,40–42 ELISA assays

can screen for specific antibodies or virus particles and are ­useful for screening a large number of samples.40–42 Recently, we developed a sapovirus ELISA that was based on hyperimmune rabbit and guinea pig antisera raised against ­sapovirus VLPs.36,38 After a number of optimization steps, the ELISA was prepared for clinical testing. Our results also showed a low sensitivity, although we found that the sapovirus ELISA was useful in detecting sapovirus GI antigens in clinical stool samples collected two days after the onset of illness. We are currently developing a new ELISA based on ­monoclonal antibodies that have broad range cross-reactivities. Several groups have designed RT-PCR primers that can detect a broad range of sapovirus strains (Table 8.1 and Figure 8.2A).19,21,23,24,43 One set of primers (sense p289 primer and antisense p290 primer) were designed to detect both norovirus and sapovirus (Table 8.1).19 In an earlier study, we detected sapovirus from stool specimens from hospitalized infants with sporadic gastroenteritis in Chiang Mai, Thailand, between July 2000 and July 2001.23 We used a single round PCR with primers (sense SV5317 primer and antisense SV5749 primer) directed against capsid gene.23 In this Thailand study, we detected three different sapovirus genotypes belonging to GI and GII, showing the usefulness of these primers. Following this study, we performed a second molecular epidemiological study using stool specimens collected from hospitalized infants with acute gastroenteritis in Thailand, between November 2002 and April 2003. However, we used a new set of nested RT-PCR primers designed by Okada et al.44 (Table 8.1). For the first PCR we used sense SV-F11 primer and antisense SV-R1 primer. For the nested PCR, we used sense SV-F21 primer and antisense SV-R2 primer. In this second Thailand study, we detected four different sapovirus genotypes, including a newly identified GV strain. A second generation set of RT-PCR primers were designed by Okada et al.45 with higher sensitivity and is currently used in our laboratory (Table 8.1). Recently, several real-time RT-PCR methods that could detect human sapovirus were developed.46–48 The advantage of real-time RT-PCR over traditional RT-PCR is that it can give a rapid result and can be used to determine the number of copies of cDNA per gram of stool sample. Two of these methods used primers and probes that targeted the capsid region, whereas the other report used primers and probes that targeted the polymerase-capsid junction. Oka et al. method was proven to detect all human sapovirus genogroups46 and is described in detail below.

8.2  Methods 8.2.1 Reagents and Equipment The essential reagents and equipment needed for sapovirus molecular detection using RT-PCR and identification by sequence analysis are listed in Table 8.2.

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Molecular Detection of Foodborne Pathogens

Table 8.1 Primers for Sapovirus Nested RT-PCR and Primers and Probes for Sapovirus Real-Time PCR Application

Target

Primer

Polarity

Location

GATTACTCCAAGTGGGACTCCAC TGACAATGTAATCATCACCATA

Sequence (5′ to 3′)

 +  –

4568a 4886a

PCR

GI and GII (other genogroups?)

p290 p289

PCR

GI and GII (other genogroups?)

SV5317 SV5749

CTCGCCACCTACRAWGCBTGGTT TGGGGVGGTASBTTTGARGYCCG

 +  –

5083b 5516b

GI, GII, GIV, GV

SV-F11 SV-R1 SV-F21 SV-R2

GCYTGGTTYATAGGTGGTAC CWGGTGAMACMCCATTKTCCAT ANTAGTGTTTGARATGGAGGG GWGGGRTCAACMCCWGGTGG

 +  –

5098b 5878b 5157b 5591b

PCR

PCR

a b c d e

SV-F13

GAYYWGGCYCTCGCYACCTAC

 + 

5074b

SV-F14 SV-R13 SV-R14 SV-F22 SV-R2

GAACAAGCTGTGGCATGCTAC GGTGANAYNCCATTKTCCAT GGTGAGMMYCCATTCTCCAT SMWAWTAGTGTTTGARATG GWGGGRTCAACMCCWGGTGG

 +  – –  +  –

5074b 5861b 5861b 5154b 5572b

GI, GII, GIV

SaV124F

GAYCASGCTCTCGCYACCTAC

 + 

5078c

GI

SaV1F

TTGGCCCTCGCCACCTAC

 + 

700d

GV GI, GII, GIV, GV GI, GII, GIV GV

SaV5F SaV1245R SaV124TP SaV5TP

TTTGAACAAGCTGTGGCATGCTAC CCCTCCATYTCAAACACTA FAM-CCRCCTATRAACCA-MGB-NQF FAM-TGCCACCAATGTACCA-MGB-NQF

 +  – – –

5112e 5163c 5105c 5142e

GI, GII, GIV, GV

Real-time PCR

 +  –

NK24 virus (AY646856). Manchester virus (X86560). Mc10 virus (AY237420). Parkville virus (U73124; partial sequence). Norwalk virus (X86560). FAM, 6-carboxyfluorescein (reporter dye); MGB, minor groove binder; NQF, nonfluorescent quencher.

A

SV-F22 (5,154)

Nested PCR

SV-F13 (5,074) SV-F14 (5,074)

B

5'

SV-R13 (5,876) SV-R14 (5,876)

first PCR

Manchester/93 genome (GI) 1

SV-R2 (5,591)

5,180

13

5,666

(5,170) RdRp

N

6,855 VP1

6,852 C 5'

7,431 A(n) 3'

C

ORF2

7,349

ORF3

Bristol/98 genome (GII) 1

14

(5,174) RdRp

N

6,853 VP1

C 6,856

ORF1

7,490 A(n) 3'

N

N-terminal of VP1

C

C-terminal of VP1

7,347

Figure 8.2  Detection of sapovirus by nested RT-PCR. The location of the primers in the RdRp/capsid junction.

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Sapoviruses

Table 8.2 Reagents and Equipment for RT-PCR and Sequencing Reagents QIAamp Viral RNA kit (Qiagen) Alcohol (80–100%) Reverse transcription reagents (e.g., SuperScript II; Invitrogen) DNA Taq polymerase reagents (e.g., Ex Taq; Takara) Random hexamers (Takara) RNase inhibitor (Toyobo) dNTPs (Roche) Terminator cycle sequence kit (e.g., BigDye; Applied Biosystems) Primers (listed in Table 8.1) Agarose Ethidium bromide Tris-acetate-EDTA (TAE) buffer DNA marker Distilled water QIAquick gel extraction kit (Qiagen)

Equipment Pipettes and tips (10 µl, 100 µl and 1000 µl) Tubes (1.5 ml, 15 ml and 50 ml) Disposable gloves Thermocycler (e.g., 2700 thermocycler; Applied Biosystems) DNA sequencer (e.g. ABI 3100 Avant sequencer; Perkin-Elmer ABI) Centrifuge (up to 10,000 × g) Heating block Vacuum manifold and pump Refrigerators (4°C and –20°C or –80°C) Vortex Measuring scale Electrophoresis tank and power supply Knife for cutting gels Beakers or flasks (600 ml) Microwave Transilluminator (an ultraviolet lightbox) Gel casting trays and combs

8.2.2 Sample Preparation A number of researchers have found that storing clinical stool samples for extended periods of time may result in RNA degradation and viral instability.19,49,50 In this regard, different research laboratories have adopted a range of different storage conditions. For example, raw stool samples, 10% stool suspensions, 10% clarified stool suspensions and extracted RNA have all been stored at such widely divergent temperatures as –20°C, –60°C, –70°C, or –80°C prior to molecular applications. In general, we keep raw stool samples at 4°C prior to diagnostic work and then store raw stool samples long term at –30°C or –80°C. Little is known about the stability of sapovirus in food or water samples, however in the case of noroviruses, samples are usually stored at 4°C or lower. Sapovirus was recently detected in untreated wastewater, treated wastewater, and a river in Japan.1,51 The methods used to concentrate the virus were different for each sample site (see below). 8.2.2.1 Primary-Treated Domestic Wastewater (Method 1)1 Viruses in 400 ml of primary-treated domestic wastewater are recovered by using the enzymatic virus elution (EVE) method.52 Primary-treated domestic wastewater is centrifuged at 9,000 × g for 15 min and the supernatant decanted. The pellet is resuspended in the EVE buffer (10 g/l of each of the following enzymes: mucopeptide N-acetylmuramoylhdrolase, carboxylesterase, chrymotrypsin, and papain) and stirred for 30 min. The suspension is centrifuged at 9,000 × g for 30 min and then the supernatant is collected and stored at –20°C until further analysis. The viruses in the supernatant are concentrated using a polyethylene glycol (PEG) precipitation method.53

PEG 6000 is added to the supernatant at a concentration of 8% and stirred at 4°C overnight. The sample is centrifuged at 9,000 × g for 90 min, and the pellet is resuspended in 4 ml of 20 mM phosphate-buffered solution (PBS; pH 8.0) and briefly agitated using a vortex. The resuspended sample is centrifuged at 9,000 × g for 10 min, and the supernatant was collected and stored at –20°C until RNA extraction. 8.2.2.2 Secondary-Treated Effluent (Method 1) and River Water (Method 1)1 Viruses in 1 l of secondary-treated effluent and river water are concentrated by PEG precipitation as follows. 100 g of PEG 6000 and 23.4 g of NaCl are added to 1 l of sample and stirred at 4°C overnight. The sample is then centrifuged at 10,000 × g for 60 min and the pellet resuspended in 4 ml of distilled water. The resuspended sample is centrifuged at 10,000 × g for 10 min and then the supernatant is collected and stored at –20°C until further analysis. The viruses in the supernatant are concentrated using the PEG precipitation method described above. Samples are stored at –20°C until RNA extraction. 8.2.2.3 Influent and Effluent Water (Method 2)2 Influent and effluent water samples are treated similarly, except that effluent samples are initially treated with sodium thiosulfate for the purpose of decholorination. Then, 2.5 M/l MgCl2 is added to the water samples to give a final concentration of 25 mM/l. 100 ml of influent and 1000 ml of effluent are passed through an HA negatively charged membrane (Nihon Millipore, Tokyo, Japan) with a 0.45-µm pore size attached to a glass filter holder. Then, 200 ml of 0.5 mmol/1 H2SO4 (pH 3) is passed through the filter (to remove magnesium ions and electropositive substances), followed by

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10 ml of 1 mmol/1 of NaOH (pH 11) (to elute the viruses). To neutralize the filtrate, 50 µl of 100 mmol/1 H2SO4 (pH 1) and 100 µl of 100 × Tris-EDTA buffer pH 8 is added. The concentrated sample is centrifuged in a Centriprep YM-50 (Nihon Millipore) according to the manufacturer’s instructions; 1,000 × g for 10 min, removal of the sample that passed through the ultrafiltration membrane and then further centrifugation at 1,000 × g for 5 min to obtain a final volume of 0.7 ml.54 Samples are stored at –20°C until RNA extraction. 8.2.2.4 River Water (Method 2)54 Another method that is used to detect sapovirus in river water was developed.1,54 5 ml of 250 mM/l AlCl3 is initially passed through an HA filter to form a cation (Al3 + )-coated filter. 500 ml of river water sample was passed through the filter, followed by successive steps using H2SO4 and NaOH as described above for influent and effluent water (method 2). The concentrated sample is centrifuged in a Centriprep YM-50 (Nihon Millipore) according to the manufacturer’s instructions to obtain a final volume of 0.7 ml and stored at –20°C until RNA extraction. 8.2.2.5 Seawater1 Viruses in 20 l of seawater are concentrated according to the method of Katayama et al.55 Seawater is filtered with an HA negatively charged membrane (Nihon Millipore) with a 0.45-µm pore size. Then, 200 ml of 0.5 mM H2SO4 is passed through the membrane to rinse out cations. Next, 10 ml of 1 mM NaOH is poured onto the membrane and the filtrate was recovered in a tube containing 0.1 ml of 50 mM H2SO4 and 0.1 ml of 100 × TE buffer. Viruses in the filtrate are further concentrated with a Centriprep Concentrator 50 system (Nihon Millipore) according to the manufacturer’s instructions to obtain a final volume of 1 ml and stored at –20°C until RNA extraction. 8.2.2.6 Oysters Unlike norovirus, human sapovirus has not yet been detected in oysters. Several methods have been developed, which are usually evaluated with a spiked virus in order to determine the efficiency of the RNA recovery. Recently, a Japanese group developed a more efficient method.56 Fresh oysters are shucked and their stomachs and digestive tracts are removed by dissection. The samples are weighed and homogenized in 10 mM PBS (pH 7.4; without magnesium or calcium) and made into a 10% suspension. The suspension is added to 0.1 ml antifoam B (Sigma, St Louis, MO) and homogenized twice at 30 sec intervals at maximum speed using an Omni-mixer (OCI Instruments, Waterbury, CT). Next, 6 ml of choloroform:butanol (1:1 vol) is added, the suspension was homogenized for 30 sec, and 170 ml Cat-Floc T (Calgon, Elwood, PA) is added. The sample is centrifuged at 3,000 × g for 30 min at 4°C and the supernatant is layered onto 1 ml of 30% sucrose solution and ultracentrifuged at 154,000 × g for 3 h at 4°C. Finally, the pellet is resuspended in 140 µl of distilled water DDW and stored at –80°C until RNA extraction.

Molecular Detection of Foodborne Pathogens

8.2.2.7 Shellfish Recently, we detected sapovirus in clam samples ­designated for human consumption.3 Clams are shucked and their stomachs and digestive diverticula are removed by dissection on the day of harvest. Then 1 g of the stomachs and digestive diverticulum (approximately 10–15 clams per package are pooled) are homogenized with an Omni-mixer in nine times their weight (approximately 9 ml) of PBS (pH 7.4; without magnesium or calcium). After centrifugation at 10,000 × g for 30 min at 4°C, all of the supernatant is layered onto 1 ml of 30% sucrose solution and ultracentrifuged at 154,000 × g for 3 h at 4°C. The supernatant is then carefully removed, and the pellet is resuspended in approximately 140 µl of distilled water and stored at –80°C until RNA extraction. 8.2.2.8 Stool Stool samples are usually diluted and clarified before RNA can be extracted using the commercial and in-house RNA extraction methods. In our laboratory, a 10% (wt/vol) stool suspension is prepared with sterilized water and centrifuged at 10,000 × g for 10 min at 4°C. However, other research laboratories have prepared the clarified supernatant in different ways. Yuen et al. prepared 10% (w/v) stool suspensions in Hanks’ complete balanced salt solution, clarified the solution by centrifuging the sample at 3,500 × g for 15 min at room temperature, followed by 7,000 × g for 30 min at 4°C.57 Drier et al. prepared 10% (wt/vol) stool suspension in PBS and clarified the solution by centrifuging the sample at 3,500 × g for 15 min.58 Houde et al. prepared 20% (wt/vol) stool suspension in PBS and clarified the solution by centrifuging the sample 2,000 × g for 3 min, followed by 16,000 × g for 5 min.59

8.2.3  Detection Procedures The rates of incidence and overall prevalence of sapovirus infections vary in each country and setting, and are likely affected by the diagnostic techniques used. Different laboratories employ different detection techniques for various reasons, including costs, equipment, time, and personnel. 8.2.3.1 EM Sapovirus was first detected by EM.9 However, this technique is tedious, since virus particles are difficult to correctly identify and the number of particles is generally low.27 To obtain a positive result by EM, approximately 106 particles per g of stool are needed as well as a skilled expert who can differentiate between a “VLP” and an actual virion. The sample preparation is straightforward although time-consuming. A 10% stool suspension is usually prepared in PBS or distilled water and then centrifuged. It is important to thoroughly vortex the suspension prior to centrifuging in order to separate the virus particles from the organic and inorganic material in the stool. The stool suspension is centrifuged at low speed (3,000 × g for 10 min) and then the supernatant collected and centrifuged at medium speed (10,000 × g for 30 min). The supernatant is then collected and centrifuged at high speed

Sapoviruses

in a 10–20% sucrose cushion (50,000 × g for 2 h) and then the pellet is resuspended in a small volume (20–50 µl) of PBS or distilled water and stored below –30°C. The method of negative staining can vary and can include (2–4%) uranyl acetate or (2–4%) phosphotungstic acid. Various kinds of EM grids can also be used, although carbon-coated grids usually provide the best resolution for sapovirus (personal communication). The virus integrity and morphology are usually stable for a certain amount of time; however, freshly prepared samples should be examined and freeze thawing may disrupt the virus morphology. The methodology for applying the sample to the grid can also vary between laboratories. In our laboratory, 10–20 µl of 1:10 sample is placed on Para film, 20 µl of water is placed on the Para film next to the sample, and then 20 µl of stain is placed on the Para film next to the water. A grid is placed in lockable forceps. The grid is touched to the sample (on the carbon side) and left to dry for 15 sec, and then the excess sample is removed from the grid with filter paper. This is repeated for the water and then the stain. Finally, the grid is left to dry for 20 min and then examined by EM at 30,000–40,000 × magnification. The resuspended pellet can also be used for molecular biology techniques, including RT-PCR and ELISA as described below. 8.2.3.2 ELISA Detection We developed an antigen ELISA detection system that was based on hyperimmune rabbit and guinea pig antisera raised against sapovirus VLPs.36 Wells of 96-well microtiter plates are coated with 100 µl of a 1:8,000 dilution of either hyperimmune rabbit (P) or preimmune rabbit (N) antisera diluted in PBS, and the plates are incubated overnight at 4°C. Wells are washed three times with PBS containing 0.1% Tween 20 (PBS-T), and then blocked with PBS containing 5% skim milk (PBS-SM) for 1 h at room temperature. After washing the wells four times with PBS-T, 100 µl of each clinical stool specimen (10% stool suspension is prepared in PBS, vortexed, centrifuged at 3,000 × g for 5 min, the supernatant is collected and further centrifuged at 10,000 × g for 30 min, and then the final supernatant collected and diluted 1:1 with PBS-T containing 1% SM [PBS-T-SM]) is added to duplicate hyperimmune rabbit and duplicate preimmune rabbit wells, and the plates are incubated for 1 h at 37°C. After washing the wells four times with PBS-T, 100 µl of a 1:8,000 dilution of hyperimmune guinea pig antiserum diluted in PBS-T-SM is added to each well, and the plates are incubated for 1 h at 37°C. Wells are washed four times with PBS-T, and then 100 µl of a 1:1,000 dilution of horseradish peroxidase (HRP)conjugated rabbit anti-guinea pig immunoglobulin G (IgG) (Cappel, West Chester, PA) diluted in PBS-T-SM is added to each well and the plates are incubated for 1 h at 37°C. Wells are washed four times with PBS-T, and then 100 µl of substrate o-phenylenediamine and H2O2 is added to each well and left in the dark for 30 min at room temperature. The reaction is stopped with the addition of 50 µl of 2 N H2SO4 to each well, and the absorbance is measured at 492 nm (A492). The cutoff value is defined as the mean plus three

107

standard deviations, and P/N ratios over the cutoff value are ­considered significantly positive.36,38 8.2.3.3 RNA Extraction RNA for RT-PCR should be free from DNA contamination, as this can generate false positives during the PCR step. Moreover, many substances found in stool samples can inhibit PCR, because the Taq polymerases require optimal conditions, including optimal pH and reagent concentrations.60 Several groups have suggested using a competitive internal control RNA to monitor inhibition of RT-PCR.61–63 Today, commercial extraction methods have replaced the earlier extraction methods and effectively remove RT-PCR inhibitors. One of the most popular methods for extracting RNA from stool samples, environmental samples or food samples is the QIAamp Viral RNA Mini Protocol (Qiagen, Hilden, Germany). The QIAamp Viral RNA Protocol is actually designed to purify viral RNA from plasma, serum, cellfree body fluids and cell-culture supernatants. However, the method can be modified to purify viral RNA from other samples as well, including stool samples and environmental samples. For a stool sample, a 10% stool suspension is prepared with sterilized water and centrifuged at 10,000 × g for 10 min at 4°C. For water samples, we use 140 µl of the concentrated water sample. For the shellfish samples, we prepare a 10% (vol/vol) of the resuspended pellet (described above). In our laboratory, this extraction method has been used to successfully detect norovirus, sapovirus, astrovirus, HEV, and Aichi virus in stool samples. It can be carried out in a standard centrifuge or a vacuum manifold, with the vacuum manifold being more useful for 24–48 samples, and the centrifuge more appropriate for a handful of samples. The entire procedure takes approximately 60–90 min (for 24 samples using a vacuum manifold). Briefly, 560 µl of 100% ethanol is added to the sample and vortexed for 15 sec. Then, 600 µl of sample is added to the QIAamp spin column and the vacuum turned on. The remaining sample (approximately 660 µl) is added to the column. After the entire sample is passed through the QIAamp spin column, the column is washed with 750 µl of buffer AW1, and then washed with 750 µl of buffer AW2. The spin column is then centrifuged at 10,000 × g for 2 min at 4°C in order to remove the residual reagents. Then, 60 µl of buffer AVE is added, the QIAamp spin column is centrifuged at 10,000 × g for 2 min at 4°C, and the RNA is collected in a clean tube. This method extracts both RNA and cellular DNA. In order to remove the DNA from the sample, it is recommended that the RNA preparation be digested with DNase, followed by heat treatment to inactivate the DNase. The RNA can be stored at –20°C or –70°C and may remain stable for up to 1 year, although we have found it to be stable for up to 2 years when stored at –80°C. 8.2.3.4 Reverse Transcription (RT) Reverse transcription (RT) can be performed using random primers, poly(T) primers or gene specific primers. A number of different reverse transcriptases are available, each with a

108

different sensitivity. Our reverse transcriptase of choice is Invitrogen Superscript III. (1) Prepare RT mixture in a final volume of 20 µl containing 10 µl of RNA in 50 ρmol random hexamer (Takara, Tokyo, Japan), 1 × Superscript III RT buffer (Invitrogen, Carlsbad, CA), 10 mM DTT (Invitrogen), 0.4 mM of each dNTP (Roche, Mannheim, Germany), 1 U RNase inhibitor (Toyobo, Osaka, Japan), and 10 U Superscript RT III (Invitrogen). (2) Perform RT at 50°C for 1 h, followed by deactivation of RT enzyme at 72°C for 15 min. Note: This standard RT method is useful for ­epidemiological studies, but a modified RT method is utilized for preparing high quantity cDNA for other molecular methods, i.e., amplification of long PCR fragments. Briefly, (i) prepare an RT mix in a PCR tube (1 × Superscript III RT buffer, 10 mM DTT, 0.4 mM of each dNTP, 1 U RNase inhibitor and 10 U Superscript RT III) and put on ice; (ii) add poly(T) reverse primer to another PCR tube with 10 µl of RNA and then add two drops of PCR oil; (iii) centrifuge the tube briefly, and put in the 94°C block for 2 min; (iv) take this tube out of the block, placed between the thumb and index finger to cool briefly (15 sec); (iv) add the RT master mix (9 µl) quickly to the bottom of this tube, centrifuge briefly and put in the 55°C block for 2–3 h; (v) deactivate the RT at 94°C for 15 min, place on ice and store the cDNA at –20°C. This cDNA is useful for producing long DNA fragments (over 3 kb) for fulllength genome analyses, expression of the capsid protein and replication studies. However, the ability to determine the fulllength norovirus genome sequence relies on the knowledge of partial sequences and degenerate primers, and on knowledge of the fact that the 5′ untranslated region (UTR) of the norovirus genome is usually conserved at the ORF2 start. 8.2.3.5 Nested PCR Numerous PCR primers that detect different regions (polymerase or capsid) have been designed and improved. Okada et al. designed a second generation of RT-PCR primers (for nested PCR) that could detect strains from all human genogroups as well as many of the different genotypes [45]. This modified primer set has also detected novel genotypes. (1) Prepare the first PCR mixture containing 5 µl of cDNA, 20 ρmol of each primer, 1 × Taq DNA polymerase buffer B, 0.2 mM of each dNTP, 2.5 U Taq polymerase, and up to 50 µl of distilled. For the first PCR, F13, F14, R13, and R14 primers are used45 (Figure 8.2 and Table 8.3). (2) Perform PCR at 94°C for 3 min followed by 35 cycles of 94°C for 30 sec, 48°C for 30 sec, 74°C for 45 sec, and a final extension of 5 min at 74°C. (3) Carry out the nested PCR using 5 µl of the first PCR in a second reaction mixture (identical to the first except for the primers) and the same PCR conditions. For the nested PCR, F22 and R2 primers are used45 (Figure 8.2 and Table 8.2).

Molecular Detection of Foodborne Pathogens

Table 8.3 Sapovirus First-Round and Nested RT-PCR Mixtures First PCR (µl)

 × _____

Second PCR (µl)

10 × ExTaq buffer

5

5

2.5 mM dNTP

4

4

SV-F13 (20 ρmol/ µl)

1

–

SV-F14 (20 ρmol/ µl)

1

–

SV-R13 (20 ρmol/ µl)

1

–

SV-R14 (20 ρmol/ µl)

1

–

SV-F22 (20 ρmol/ µl)

–

1

SV-R2 (20 ρmol/ µl)

–

1

cDNA or 1st PCR product ExTaq DNA polymerase

5

–

2

  0.25

  0.25

Distilled water

31.75

36.75

Total per tube

50

50

 × _____

–

Note: For molecular methods that require high fidelity, we used KOD DNA polymerase, which has a unique proofreading ability and is faster and more accurate than conventional DNA polymerases. The proofreading ability results in a lower PCR mutation frequency and considerably higher elongation rates than those achieved by conventional DNA polymerases. 8.2.3.6 Real-time RT-PCR The real-time RT-PCR method presented below was reported recently for detection of all human sapovirus genogroups.46 (1) Extract RNA with the Qiagen method. (2) Add viral RNA (10 µl) to a reaction mixture (5 µl) containing DNase I buffer (150 mM Tris-HCl, pH 8.3, 225 mM KCl, 9 mM MgCl2) and 1 U of RQ1; incubate the reaction mixture first at 37°C for 30 min to digest DNA and then at 75°C for 5 min to inactivate the enzyme. (3) Add DNase I-treated RNA (15 µl) to 15 µl of another mixture containing RT buffer, 1 mM of each dNTP, 10 mM dithiothreitol, 50 pmol of random hexamers, 30 units of RNase OUT and 200 units of SuperScript III RNaseH (–) reverse transcriptase DNase; perform RT at 37°C for 15 min followed by 50°C for 1 h, and store the solution at –20°C. (4) Prepare quantitative real-time RT-PCR in a 25 µl of a reaction volume using a QuantiTect Probe PCR Kit containing 2.5 µl of cDNA, 12.5 µl of Quantitect Probe PCR Master Mix, 400 nM of each primer (SaV124F, SaV1F, SaV5F, and SaV1245R), and 5 ρmol of TaqMan MGB probes (SaV124TP and SaV5TP) (Table 8.4). The four primers (SaV124F, SaV1F, SaV5F, and SaV1245R) and two TaqMan MGB probes (SaV124TP and SaV5TP) are designed

109

Sapoviruses

8.3 Conclusions and Future Perspectives

Table 8.4 Sapovirus Real-Time PCR Mixture µl 2 × real-time PCR Master mix

12.5

SaV124F (10 ρmol/ µl)

1

SaV1F (10 ρmol/ µl)

1

SaV5F (10 ρmol/ µl)

1

SaV1245R (10 ρmol/ µl)

1

SaV124TP (10 ρmol/ µl)

1

SaV5TP (10 ρmol/ µl)

1

cDNA or plasmid

 × _____

  2.5

Distilled water

5

Total per well

25

–

(using multiple alignment analysis of 27 sapovirus sequences) to hybridize against the highly conserved nucleotides between 5078 and 5181 with respect to Mc10 virus, and used here to detect sapovirus GI, GII, GIV, and GV sequences in a single reaction tube. (5) Perform PCR amplification with a 7500 Fast RealTime PCR System (Applied Biosystems, Foster City, CA) under the following conditions: initial denaturation at 95°C for 15 min to activate DNA polymerase, followed by 40 cycles of amplification with denaturation at 94°C for 15 sec, and annealing and extension at 62°C for 1 min. (6) Collect and analyze amplification data with Sequence Detector software version 1.3 (Applied Biosystems); use a ten-fold serial dilution of standard cDNA plasmid (2.5 × 107–2.5 × 101 copies) to quantify the viral copy numbers in reaction tubes. The detection limit of the real-time RT-PCR is approximately 1.3 × 105 copies of sapovirus RNA per gram of stool sample. 8.2.3.7  Phylogenetic Analysis (1) Excise the PCR-generated amplicons from the gel and purify by the QIAquick gel extraction kit (Qiagen). (2) Sequence the PCR products directly with the terminator cycle sequence kit (version 3.1) and determined with the ABI 3100 Avant sequencer (Perkin-Elmer ABI, Boston, MA); or (3) Clone the PCR products into plasmid vector (e.g., pCR2.1; Invitrogen) in case foodborne outbreaks due to multiple strains are suspected. Four or more colonies should be sequenced. (4) Align nucleotide sequences using Clustal X and calculate the distances by Kimura’s two-parameter method.64 Generate phylogenetic trees with bootstrap analysis from 1000 replicas by the neighborjoining method as described previously.64

New methods and technologies are continuously been developed and tested. More rapid, reliable, and sensitive detection systems will benefit diagnostic laboratories and may improve outbreak management. Methods that can reduce the time needed for complete analysis will allow for more samples to be processed, which in turn will provide better information on these viruses. Reliable and highly sensitive methods that can be used worldwide, will improve surveillance and reporting. A number of companies have developed kits that can extract RNA from up to 96 samples at one time. Ambion developed a rapid high throughput (96 samples) viral RNA isolation kit (MagMax-96 Viral RNA Isolation Kit; Ambion, Austin, TX) that captures both viral RNA and DNA onto microspherical paramagnetic beads. Promega developed an automated system that can extract total RNA from 96 samples (SV 96 Total RNA Isolation System; Promega, Madison, WI) and capture the extracted RNA onto glass fibers. Beckman CoulterQiagen developed at kit that can extract DNA and RNA (96 samples) from plasma, serum, and other cell-free body fluids (QIAamp Virus BioRobot 9604 Kit; Qiagen) and capture the nucleic acid onto a silica-gel membrane. Recently other detection methods have been developed for norovirus, including isothermal nucleic acid sequencebased amplification (NASBA), real-time RT-loop mediated isothermal amplification (LAMP) and real-time LightCycler RT-PCR.65–67 Immunochromatography (IC) is another rapid detection method, and a method to detect norovirus by IC was recently developed by a company in Japan (Immuno-Probe Company, Saitama, Japan). Recently, a microarray assay was also developed for the detection and typing of noroviruses.68 These methods have yet to be developed for sapovirus, but since the number of sapovirus-associated outbreaks of gastroenteritis appears to be steadily increasing, more rapid and reliable methods will be sought after.

References

1. Hansman, G.S. et al. Sapovirus in water, Japan. Emerg. Infect. Dis., 13, 133, 2007. 2. Haramoto, E. et al. Quantitative detection of sapoviruses in wastewater and river water in Japan. Lett. Appl. Microbiol., 46, 408, 2008. 3. Hansman, G.S. et al. Human sapovirus in clams, Japan. Emerg. Infect. Dis., 13, 620, 2007. 4. Costantini, V. et al. Human and animal enteric caliciviruses in oysters from different coastal regions of the United States. Appl. Environ. Microbiol., 72, 1800, 2006. 5. Hansman, G.S. et al. An outbreak of gastroenteritis due to Sapovirus. J. Clin. Microbiol., 45, 1347, 2007. 6. Hansman, G.S. et al. Recombinant Sapovirus gastroenteritis, Japan. Emerg. Infect. Dis., 13, 786, 2007. 7. Johansson, P.J. et al. A nosocomial sapovirus-associated outbreak of gastroenteritis in adults. Scand. J. Infect. Dis., 37, 200, 2005. 8. Noel, J.S. et al. Parkville virus: a novel genetic variant of human calicivirus in the Sapporo virus clade, associated with an outbreak of gastroenteritis in adults. J. Med. Virol., 52, 173, 1997.

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9. Chiba, S. et al. An outbreak of gastroenteritis associated with calicivirus in an infant home. J. Med. Virol., 4, 249, 1979. 10. Katayama, K. et al. Novel recombinant sapovirus. Emerg. Infect. Dis., 10, 1874, 2004. 11. Hansman, G.S. et al. Intergenogroup recombination in sapoviruses. Emerg. Infect. Dis., 11, 1916, 2005. 12. Lole, K.S. et al. Full-length human immunodeficiency virus type 1 genomes from subtype C-infected seroconverters in India, with evidence of intersubtype recombination. J. Virol., 73, 152, 1999. 13. Sakuma, Y. et al. Prevalence of antibody to human calicivirus in general population of northern Japan. J. Med. Virol., 7, 221, 1981. 14. Grohmann, G. et al. Outbreak of human calicivirus gastroenteritis in a day-care center in Sydney, Australia. J. Clin. Microbiol., 29, 544, 1991. 15. Hedlund, K.O. et al. Epidemiology of calicivirus infections in Sweden, 1994-1998. J. Infect. Dis., 181 (Suppl 2), S275, 2000. 16. Schuffenecker, I. et al. Genetic classification of “Sapporo-like viruses”. Arch. Virol., 146, 2115, 2001. 17. Nakata, S. et al. Members of the family caliciviridae (Norwalk virus and Sapporo virus) are the most prevalent cause of gastroenteritis outbreaks among infants in Japan. J. Infect. Dis., 181, 2029, 2000. 18. Robinson, S. et al. Epidemiology of human Sapporo-like caliciviruses in the South West of England: molecular characterisation of a genetically distinct isolate. J. Med. Virol., 67, 282, 2002. 19. Jiang, X. et al. Design and evaluation of a primer pair that detects both Norwalk- and Sapporo-like caliciviruses by RT-PCR. J. Virol. Methods, 83, 145, 1999. 20. Wolfaardt, M. et al. Incidence of human calicivirus and rotavirus infection in patients with gastroenteritis in South Africa. J. Med. Virol., 51, 290, 1997. 21. Vinje, J. et al. Molecular detection and epidemiology of Sapporo-like viruses. J. Clin. Microbiol., 38, 530, 2000. 22. Buesa, J. et al. Molecular epidemiology of caliciviruses causing outbreaks and sporadic cases of acute gastroenteritis in Spain. J. Clin. Microbiol., 40, 2854, 2002. 23. Hansman, G.S. et al. Genetic diversity of norovirus and sapovirus in hospitalized infants with sporadic cases of acute gastroenteritis in Chiang Mai, Thailand. J. Clin. Microbiol., 42, 1305, 2004. 24. Okada, M. et al. Molecular epidemiology and phylogenetic analysis of Sapporo-like viruses. Arch. Virol., 147, 1445, 2002. 25. Farkas, T. et al. Prevalence and genetic diversity of human caliciviruses (HuCVs) in Mexican children. J. Med. Virol., 62, 217, 2000. 26. Pang, X.L. et al. Human caliciviruses in acute gastroenteritis of young children in the community. J. Infect. Dis., 181 Suppl 2, S288, 2000. 27. Matson, D.O. et al. Human calicivirus-associated diarrhea in children attending day care centers. J. Infect. Dis., 159, 71, 1989. 28. Matson, D.O. et al. Asymptomatic human calicivirus infection in a day care center. Pediatr. Infect. Dis. J., 9, 190, 1990. 29. Lopman, B.A. et al. Viral gastroenteritis outbreaks in Europe, 1995–2000. Emerg. Infect. Dis., 9, 90, 2003. 30. Guo, M. et al. Expression and self-assembly in baculovirus of porcine enteric calicivirus capsids into virus-like particles and their use in an enzyme-linked immunosorbent assay for antibody detection in swine. J. Clin. Microbiol., 39, 1487, 2001.

Molecular Detection of Foodborne Pathogens 31. Chen, R. et al. Inter- and intragenus structural variations in caliciviruses and their functional implications. J. Virol., 78, 6469, 2004. 32. Jiang, X. et al. Expression and characterization of Sapporolike human calicivirus capsid proteins in baculovirus. J. Virol. Methods, 78, 81, 1999. 33. Numata, K. et al. Molecular characterization of morphologically typical human calicivirus Sapporo. Arch. Virol., 142, 1537, 1997. 34. Oka, T. et al. Expression of sapovirus virus-like particles in mammalian cells. Arch. Virol., 151, 399, 2005. 35. Hansman, G.S. et al. Antigenic diversity of human Sapoviruses. Emerg. Infect. Dis., 13, 1519, 2007. 36. Hansman, G.S. et al. Development of an antigen ELISA to detect sapovirus in clinical stool specimens. Arch. Virol., 151, 551, 2006. 37. Hansman, G.S. et al. Cross-reactivity among sapovirus recombinant capsid proteins. Arch. Virol., 150, 21, 2005. 38. Hansman, G.S. et al. Characterization of polyclonal antibodies raised against sapovirus genogroup five virus-like particles. Arch. Virol., 150, 1433, 2005. 39. Hansman, G.S. et al. Antigenic diversity of human sapoviruses. Emerg. Infect. Dis., 13, 1519, 2007. 40. Nakata, S. et al. Microtiter solid-phase radioimmunoassay for detection of human calicivirus in stools. J. Clin. Microbiol., 17, 198, 1983. 41. Chiba, S. et al. Sapporo virus: history and recent findings. J. Infect. Dis., 181 Suppl 2, S303, 2000. 42. Nakata, S. et al. Detection of human calicivirus antigen and antibody by enzyme-linked immunosorbent assays. J. Clin. Microbiol., 26, 2001, 1988. 43. Okada, M. et al. The detection of human sapoviruses with universal and genogroup-specific primers. Arch. Virol., 151, 2503, 2006. 44. Okada, M. et al. Molecular epidemiology and phylogenetic analysis of Sapporo-like viruses. Arch. Virol., 147, 1445, 2002. 45. Hansman, G.S. et al. Genetic diversity of Sapovirus in children, Australia. Emerg. Infect. Dis., 12, 141, 2006. 46. Oka, T. et al. Detection of human sapovirus by real-time reverse transcription-polymerase chain reaction. J. Med. Virol., 78, 1347, 2006. 47. Gunson, R. N. et al. The real-time detection of sapovirus. J. Clin. Virol., 35, 321, 2006. 48. Chan, M. C. et al. Sapovirus detection by quantitative realtime RT-PCR in clinical stool specimens. J. Virol. Methods, 134, 146, 2006. 49. Vinje, J. et al. International collaborative study to compare reverse transcriptase PCR assays for detection and genotyping of noroviruses. J. Clin. Microbiol., 41, 1423, 2003. 50. Trujillo, A.A. et al. Use of TaqMan real-time reverse transcription-PCR for rapid detection, quantification, and typing of norovirus. J. Clin. Microbiol., 44, 1405, 2006. 51. Ueki, Y. et al. Norovirus pathway in water environment estimated by genetic analysis of strains from patients of gastroenteritis, sewage, treated wastewater, river water and oysters. Water Res., 39, 4271, 2005. 52. Sano, D. et al. Detection of enteric viruses in municipal sewage sludge by a combination of the enzymatic virus elution method and RT-PCR. Water Res., 37, 3490, 2003. 53. Lewis, G.D. and Metcalf, T.G. Polyethylene glycol precipitation for recovery of pathogenic viruses, including ­hepatitis A virus and human rotavirus, from oyster, water, and sediment samples. Appl. Environ. Microbiol., 54, 1983, 1988.

Sapoviruses 54. Haramoto, E. et al. Application of cation-coated filter method to detection of noroviruses, enteroviruses, adenoviruses, and torque teno viruses in the Tamagawa River in Japan. Appl. Environ. Microbiol., 71, 2403, 2005. 55. Katayama, H. et al. Development of a virus concentration method and its application to detection of enterovirus and norwalk virus from coastal seawater. Appl. Environ. Microbiol., 68, 1033, 2002. 56. Nishida, T. et al. Genotyping and quantitation of noroviruses in oysters from two distinct sea areas in Japan. Microbiol. Immunol., 51, 177, 2007. 57. Yuen, L. K. et al. Heminested multiplex reverse transcriptionPCR for detection and differentiation of Norwalk-like virus genogroups 1 and 2 in fecal samples. J. Clin. Microbiol., 39, 2690, 2001. 58. Dreier, J. et al. Enhanced reverse transcription-PCR assay for detection of norovirus genogroup I. J. Clin. Microbiol., 44, 2714, 2006. 59. Houde, A. et al. Comparative evaluation of RT-PCR, nucleic acid sequence-based amplification (NASBA) and real-time RT-PCR for detection of noroviruses in faecal material. J. Virol. Methods, 135, 163, 2006. 60. Hale, A.D. et al. Comparison of four RNA extraction methods for the detection of small round structured viruses in faecal specimens. J. Virol. Methods, 57, 195, 1996.

111 61. Atmar, R.L. and Estes, M.K. Diagnosis of noncultivatable ­gastroenteritis viruses, the human caliciviruses. Clin. Microbiol. Rev., 14, 15, 2001. 62. Wang, Q.H. et al. Development of a new microwell hybridization assay and an internal control RNA for the detection of porcine noroviruses and sapoviruses by reverse transcriptionPCR. J. Virol. Methods, 132, 135, 2006. 63. Schwab, K.J. et al. Use of heat release and an internal RNA standard control in reverse transcription-PCR detection of Norwalk virus from stool samples. J. Clin. Microbiol., 35, 511, 1997. 64. Katayama, K. et al. Phylogenetic analysis of the complete genome of 18 Norwalk-like viruses. Virology, 299, 225, 2002. 65. Fukuda, S. et al. Rapid detection of norovirus from fecal specimens by real-time reverse transcription-loop-mediated isothermal amplification assay. J. Clin. Microbiol., 44, 1376, 2006. 66. Greene, S.R. et al. Evaluation of the NucliSens Basic Kit assay for detection of Norwalk virus RNA in stool specimens. J. Virol. Methods, 108, 123, 2003. 67. Pang, X. et al. Evaluation and validation of real-time reverse transcription-pcr assay using the LightCycler system for detection and quantitation of norovirus. J. Clin. Microbiol., 42, 4679, 2004. 68. Jaaskelainen, A.J. and Maunula, L. Applicability of microarray technique for the detection of noro- and astroviruses. J. Virol. Methods, 136, 210, 2006.

9 Slow Viral Diseases Takashi Onodera, Guangai Xue University of Tokyo

Akikazu Sakudo Osaka University

Gianluigi Zanusso University of Verona

Katsuaki Sugiura

Food and Agricultural Materials Inspection Center

Contents 9.1 Introduction........................................................................................................................................................................113 9.1.1 Definition and Classification of Slow Viral Diseases.............................................................................................113 9.1.2 Biology, Pathogenesis, and Medical Importance....................................................................................................114 9.1.3 Laboratory Diagnosis of Slow Viral Diseases........................................................................................................116 9.2 Methods............................................................................................................................................................................. 120 9.2.1 Reagents and Equipment........................................................................................................................................ 120 9.2.2 Sample Collection and Preparation....................................................................................................................... 120 9.2.3 Detection Procedures............................................................................................................................................. 120 9.3 Conclusions and Future Perspectives................................................................................................................................ 122 References.................................................................................................................................................................................. 123

9.1  Introduction 9.1.1 Definition and Classification of Slow Viral Diseases Prion diseases or transmissible spongiform encephalopathies (TSEs) are fatal neurological disorders that include Creutzfeldt–Jakob disease (CJD) in humans, scrapie in sheep and goats, bovine spongiform encephalopathy (BSE) in cattle, and chronic wasting disease (CWD) in cervids. A key event in prion diseases is the conversion of the cellular, hostencoded prion protein (PrPC) to its abnormal isoform (PrPSc) predominantly in the central nervous system of the infected host.1 There is increasing evidence that the major and possibly only component of the infectious agent is PrPSc or a prion protein (PrP) folding intermediate.2 PrPC is a cell surfaceanchored glycoprotein whose function is not well characterized.3 PrPSc is derived from PrPC in a post-translational process that appears to involve PrPC–PrPSc molecular interactions.4 The crucial role of PrPC expression in prion infection and PrPSc formation has been demonstrated in transgenic mice with an ablated PrP murine gene.5 Transgenic animals have since been used extensively to unravel the influence

of specific PrP amino acid residues or domains on prion susceptibility.6–11 Kuru and the transmissible agents in dementias have been classified in a group of virus-induced slow infections that we have described as sub-acute spongiform virus encephalopathies because of the strikingly similar histopathological lesions they induce (Table 9.1). Slow virus diseases that we have been variously designated as sub-acute spongiform encephalopathies, transmissible amyloidoses of the brain, or transmissible cerebral amyloidoses (TCA) will be discussed individually below. Kuru was the first chronic or sub-acute degenerative disease in humans that was shown to be provoked by a slow virus infection, and as such has attracted worldwide attention and stimulated the search for similar virus infections as the possible cause of other sub-acute and chronic diseases in humans.12–17 Such a slow progressive infection in humans, with an incubation period of many years and an absence of any inflammatory responses to the slow destruction of brain cells, was previously unknown. In veterinary medicine, Sigurdsson17 had shown such slow virus infection in sheep. The elucidation of the etiology of kuru has led to the discovery that worldwide pre-senile dementias, CJD and its variants, with basically 113

114

Table 9.1 Transmissible Spongiform Encephalopathies with Mutations Responsible for Inherited Genetic forms of the Disease In humans Kuru Creutzfeldt–Jakob disease Sporadic Iatrogeric (human growth hormone, pituitary gonadotropin, dura mater and corneal transplants, stereotactic electrodes) Familial (178asn, 200lys, 210ile, octapeptide inserts 2, 4–9) Variant Gerstmann–Sträussler–Scheinker syndrome (GSS is only genetic. Mutations are classified according to disease phenotype) Spastic paraparesis with dementia (105leu) Familial, Alzheimer-like (145stop, 198ser, 217arg) Atypical dementia (octapeptide repeat insertions 6, 7) Fatal familial insomnia (178asn) In animals Scrapie (sheep, goats) Transmissible mink encephalopathy Chronic wasting disease (mule, deer, elk) Bovine spongiform encephalopathy Exotic ungulate spongiform encephalopathy (nyala, gemsbok, Arabian oryx, greater kudu, eland, moufflon) Feline spongiform encephalopathy (cats, albino tigers, puma, cheetah)

Molecular Detection of Foodborne Pathogens

Table 9.2 Conventional Virus-Induced Slow Infections of Man Virus

Picornaviridae Poliovirus Echovirus Paramyxoviridae Measles Togaviridae Rubella Flaviviridae Tick-borne encephalitis (RSSE) Hepatitis C

Diseases RNA viruses Chronic meningoencephalitis In immunodeficient patients Sub-acute postmeasles leuco-encephalitis Sub-acute sclerosing panencephalitis Progressive congenital rubella Progressive panencephalitis Epilepsia partialis continua (Kozhevnikov’s epilepsy) and progressive bulbar palsy in Russia Chronic hepatitis C (non-A, non-B hepatitis)

Retroviridae Lentivirus HIV-1, HIV-2 HIV neuromyeloencephalopathy Oncovirus HTLV-1, HTLV-2 Adult T-cell leukemia and mycosis fungoides, HAM/TSP neuromyeloencephalopathy Jamaican neuropathy, tropical spastic paraparesis Rhabdoviridae Rabies lyssavirus Rabies encephalitis DNA viruses

similar cellular lesions, are also transmissible and caused by a very similar, unconventional “virus” or prion.18 In all of TCA in humans, infectious amyloids are formed from the TCA amyloid precursor protein (PrP) specified on the short arm of chromosome 20 in human and chromosome 2 in mouse. There are other slow infections of the CNS in humans that are caused by conventional viruses, including measles virus, papovaviruses (JCV and SV40-PML), rubella virus, cytomegalovirus, herpes simplex virus, adenovirus types 7 and 32, Russian spring–summer encephalitis (RSSE) viruses, and the human retroviruses (human T-cell lymphotropic viruses-I: HTLV1, and human immunodeficiency viruses: HIV) (Table 9.2). However, unlike these conventional viruses, the “unconventional viruses” of the sub-acute spongiform encephalopathies are truly slow in their replication, with long doubling times. Moreover, these alone appear to be viruses that have made it necessary to alter our conceptions of the possible range of virus structure. The process of infection appears to be a seeding by a “virus,” which is a nucleating agents inducing and then automatically accelerating the conformational transition in the amyloid submit protein. In that process host-specified precursor protein (PrPC) is converted to an insoluble cross-β-plated configuration (PrPSc).19–21 Oligomers of microfilaments of this amyloid submit protein nucleate its own polymerization, crystallization, and precipitation as insoluble arrays of amyloid fibrils. Thus, proteolytic cleavage and conformational change of the precursor and oligomeric

Adenoviridae Adenovirus 7 and 32 Papovaridae JC virus SV40 Hepadnoviridae Hepatitis B Herpesviridae Herpes Simplex Cytomegalovirus Epstein-Barr virus Varicella-Zoster virus Herpes saimiri (B-virus)

Sub-acute encephalitis Progressive multifocal leucoencephalopathy Progressive multifocal leucoencephalopathy Homologous serum jaundice Sub-acute encephalitis Cytomegalovirus brain infection Chronic infectious mononucleosis Herpes zoster (sub-acute encephalitis) Chronic herpes B encephalitis

assembly of this structurally altered polypeptide produces a fibril amyloid enhancing factor22 with apparent infectious properties (Table 9.2).

9.1.2  Biology, Pathogenesis, and Medical Importance Several features distinguish prions from viruses. First, prions can exist in multiple forms, whereas viruses exist in a single form with distinct ultrastructural morphology. Second, prions are non-immunogenic in contrast to viruses, which almost always provoke an immune response. Third, there is no evidence for an essential nucleic acid within the infectious prion particle, whereas viruses have a nucleic acid genome that serves as the template for the synthesis of viral progeny. Forth, the only known component of the prion is PrPSc, which

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is encoded by a chromosomal gene, whereas viruses are composed of nucleic acid, proteins, and often other constituents. PrPSc is derived from PrPC by a post-translational process. The molecular events in the conversion are unknown but may involve only a change in the conformation of the protein. PrPSc may be distinguished from PrPC by its different biochemical and biophysical properties.22–26 Limited proteolysis of PrPSc produces a smaller, protease-resistant molecule of about 140 amino acids, designated PrP.27–30 Under the same conditions PrPC is completely protease-digested. The amino acid sequence of PrPSc that has been established by protein sequencing and mass spectrometry is identical to that deduced from the genomic DNA sequence. No proteins other than PrPSc have been consistently found in fractions enriched for prion infectivity. Prion diseases are a group of neurodegenerative disorders affecting mammals (Table 9.1). The diseases are transmissible under some circumstances, but unlike other transmissible disorders prion diseases they can also be caused by mutations in the host PrP gene. The mechanism of prion spread among sheep and goats developing natural scrapie is unknown. CWD, transmissible mink encephalopathy (TME), BSE, feline spongiform encephalopathy (FSE), and exotic ungulate encephalopathy (EUE) are all thought to occur after the consumption of prion-infected materials. Similarly, kuru of New-Guinea Fore people is thought to have resulted from the consumption of brain tissue during ritualistic cannibalism. Familial CJD, Gerstmann–Sträussler–Scheinker (GSS), and Fatal familial insomnia (FFI) are all dominant, inherited prion diseases have been show to be genetically linked to mutations in the PrP gene, while iatrogenic CJD cases can be traced to inoculation of prions through human pituitary derived growth hormone, cornea transplants, dura mater grafts, or cerebral electrode implants; although the number of cases recorded to date is small. Most cases of CJD are sporadic, probably the result of somatic mutation of the PrP gene or the spontaneous conversion of PrPC into PrPSc. About 10–15% of CJD cases and virtually all cases of GSS and FFI appear to be caused by germline mutations in the PrP gene. Twelve different PrP genes have been shown to segregate with human prion diseases (Table 9.3 and Figure 9.1). Variant CJD (vCJD) was identified by the CJD Surveillance Unit in Great Britain,27 followed by a report of a similar case in France.28 The patients in these reports differed from typical cases of sporadic CJD in terms of the relatively young age at disease onset (average 26.3 years), the prolonged duration of illness (average 14 months), and the clinical features with psychiatric symptoms, then with ataxia and the pain at the extremities, with myoclonus or chorea occurring late in the illness. Dementia was not evident until the final stages of the illness, when it was often accompanied by cortical blindness and akinetic mutism (Table 9.4). None of these patients showed changes in electroencephalogram readings that are characteristic of sporadic CJD, and all were methionine homozygous at codon 129 with no pathogenic mutations in the human prion protein gene (PRNP).

Table 9.3 Prion Diseases and Gene Mutations Disease PrP

Gene Mutation

Gerstmann–Sträussler–Scheinker syndrome Gerstmann–Sträussler–Scheinker syndrome Gerstmann–Sträussler–Scheinker syndrome Gerstmann–Sträussler–Scheinker syndrome Familial Creutzfeldt–Jakob disease

(PrP P102L) (PrP P105L, 129V) (PrP A117V) (PrP G131V, 129M)

Fatal Familial Insomnia Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease Gerstmann–Sträussler–Scheinker syndrome Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease Gerstmann–Sträussler–Scheinker syndrome Gerstmann–Sträussler–Scheinker syndrome Familial Creutzfeldt–Jakob disease Familial Creutzfeldt–Jakob disease or Gerstmann–Sträussler–Scheinker syndrome

(PrP D178N, 129M) (PrP D178N, 129V) (PrP V180I, 129M) (PrP T183A, 129M) (PrP T188A, 129M) (PrP E196K) (PrP F198S, 129V) (PrP E200K, 129 M or V) (PrP V203I, 129M) (PrP R208H, 129M) (PrP V210M, 129M) (PrP E211Q, 129M) (PrP Q212P, 129M) (PrP Q217R, 129V) (PrP M232R, 129M) (PrP octapeptide repeat insertions)

The neuropathological features in these cases were remarkably similar and presented a spectrum of changes that were highly unusual for sporadic CJD. Although spongiform changes were present in all cases in the cerebral cortex, the most striking abnormality was the presence of multiple kurutype PrP plaques, many of which were surrounded by a halo of spongiform change. Similar plaques were present in other regions of the cerebrum and cerebellum. Spongiform change was most evident in the basal ganglia and thalamus, with severe thalamic astrocytosis. Immunocytochemistry showed widespread accumulation of PrP in the brain, particularly in the cerebellar cortex and occipital cortex. The relationship of these cases to sporadic CJD in older patients is unclear and further studies are required to characterize this unusual phenotype and investigate the possibility that these cases are usually related to the BSE agent via the human food chain. Epidemiological studies have not shown increased risk in particular occupations that may be exposed to human or animal prions, although individual CJD cases in two histopathology technicians, a neuropathologist, and a neurosurgeon have been documented. While there have been concerns that CJD may be transmitted by blood transfusion, extensive epidemiological analysis in the UK has found that the frequency of blood transfusion and donation was not different over 200 cases of CJD and a matched control population.29 Recent reports show that vCJD may be transmitted by blood. One case of probable transfusion-transmitted vCJD infection has been reported,

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Molecular Detection of Foodborne Pathogens

A

B

C

6C4

D

E

F

T2

Type-1 CJD

Type-2 CJD

Figure 9.1  Immunohistochemical staining with monoclonal antibodies in human brain tissue with CJD. Type-1 (synaptic type) CJD specimen were intensively stained with mAbs 6C4 (Panel A) and T2 (Panel D) in the molecular layer of the cerebellum, with findings showing granular (Type-1; synapsis type) staining. Type-2 (Kuru type) spherical plaques (Panel B, C, E, F) in the cerebellum from a CJD patient with Met/Val polymorphism at codon 129 were differently stained with mAbs 6C4 and T2. The former was characterized by negative staining in the core of plaques, while the latter portrayed homogeneous staining of the central area (Panel C, F). (From Hosokawa, T., et al., Microbiol. Immunol., 52, 25, 2008.)

Table 9.4 Diagnostic Criteria for vCJD I

Progressive neuropsychiatric disorder Duration of illness > 6 months Routine investigations do not suggest an alternative diagnosis No history of potential iatrogenic exposure

II

Early psychiatric symptoms Persistent painful sensory symptoms Ataxia Mycolonus or chorea or dystonia Dementia

III

IV

Electroencephalogram does not show the typical appearance of sporadic CJD (or no electroencephalogram done) Bilateral pulvnar high signal on MRI scan Positive tonsil biopsy

and one case of subclinical infection has been detected.30,31 On February 9, 2006, a third case was announced by the UK Health Protection Agency. Each of the three patients had received a blood transfusion from a donor who subsequently developed clinical vCJD, which indicates that transfusion caused the infection. However, a policy to exclude potential donors who had received a fusion would not have prevented at least the first two cases because the corresponding donors had not received any blood transfusion. Diagnostic tools to

detect prions in blood are under development,32 but no routine test for the presence of the infectious agents of vCJD is available at present. Therefore, questions arise as to whether an infection like vCJD could spread endemically through blood donation alone and to what extent the exclusion of potential donors with a history of transfusion would influence the transmission of such an infection (i.e., how many deaths due to the infection could be prevented?). Transmission of CJD to rodents following intracerebral inoculation with buffy coat from CJD cases has been ­reported.33 The transmission study on rodents or chimpanzee have been done with blood from sporadic or genetic CJD, indicating that the transmissibility of sporadic CJD do not occur by transfusion.34 In addition, differently from vCJD this transmitted strain is not lymphotrophic.

9.1.3 Laboratory Diagnosis of Slow Viral Diseases A specific diagnosis of prion diseases relies on the detection of PrPSc (Table 9.5). However, the diagnostic methods currently used such as immunohistochemistry, western blotting, and enzyme-linked immunosorbent assay (ELISA) techniques apparently lack sufficient specificity to detect PrPSc in cerebrospinal fluid although transmission studies indicate the presence of some infectivity in some species.35 PrPSc can be distinguished from PrPC by its high beta-sheet content,36 its partial resistance to protease digestion,37 and its tendency to form large aggregates both in vivo and in vitro.38 The formation of multimeric aggregates can be assumed to be closely

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Slow Viral Diseases

Table 9.5 Diagnostic Methods for Prion Infections Method

Principle

Western blotting ELISA

PK-resistant PrP PK-resistant PrP

Immunohistochemistry Bioassay Cell culture assay Histoblot

PK-resistant PrP PK-resistant PrP, incubation time or infectivity titer PK-resistant PrP or infectivity titer PK-resistant PrP

Cell blot

PK-resistant PrP

Slot blot

PK-resistant PrP

PET blot

PK-resistant PrP

PMCA

PK-resistant PrP

CDI DELFIA

PrP conformation Insoluble PrP

Capillary gel electrophoresis

PK-resistant PrP

FCS

Aggregation of PrP

Aptamer

PrP conformation

FT-IR spectroscopy MUFS Flow microbead immunoassay Surrogate marker

Alterations of spectral feature Alterations of spectral feature PK-resistant PrP Change of expression level of 14-3-3 protein, erythroid-specific marker or plasminogen

Procedural Detail Detect PK-resistant PrP on the membrane Detect PK-resistant PrP adsorbed on microtiter plates by anti-PrP antibody Immunostain of tissue sections Transmission to mice Transmission to cells Cryosection is blotted onto membrane before PK-treatment and immunolabeling with anti-PrP antibody Grow the cells on cover slip, directly transfered to membrane, and detect the PK-resistant PrP using anti-PrP antibody. Filter the cell lysate through nitrocellulose membrane, and detect the PK-resistant PrP using anti-PrP antibody. Paraffin-embedded tissue section is collected on membrane, and PK-resistnt PrP is immunolabeled with anti-PrP antibody Amplfication of misfolding protein by cycles of incubation and sonication Specific antibody binding to denatured and native forms of PrP Measure a percentage of the insoluble PrP extracted by two concentrations of guanidine hydrochloride Competition between fluorescein labeled synthetic PrP peptide and PrP present in samples is assayed by separation of free and antibodypeptide peaks using capillary electrophoresis PrP aggregates were labeled by anti-PrP antibody tagged with fluorecent dyes, resulting in intensity fluorescent target, which were measured by dual-color fluorescence intensity distribution analysis Use RNA aptamers that specifically recognize PrPC and/or PrPSc conformation. Analyze FT-IR spectra with statistical analysis Analyze spectra of emission excited by ultraviolet radiation Detect PK-resistant PrP using flowcytometer with anti-PrP antibody coupled with microbeads Detect the change by two-demensional gel electrophoresis, differential display reverse-transcriptase PCR or Western blotting of surrogate marker proteins for prion diseases

References [108] [109] [110] [111] [112] [78] [54] [56] [43,77] [81] [57] [113] [78,88]

[41,91]

[104] [92] [93] [79] [114–117]

Source: Modified from Sakudo, A. et al., J. Vet. Med. Sci., 69, 329, 2007. With permission from The Japanese Society of Veterinary Science. Notes: PET: Paraffin-embedded tissue, PMCA: Protein misfolding cyclic amplification, CDI: Conformation-dependent immunoassay, DELFIA: Dissociationenhanced lanthanide fluorescent immunoassay, FCS: Fluorescence correlation spectroscopy, FT-IR: Fourier transformed infrared, ELISA: Enzymelinked immunosorbent assay, MUFS: Multi-spectral ultraviolet fluorescence spectroscopy, PCR: Polymerase chain reaction, PK: Proteinase K, PrP: Prion protein, PrPC: Cellular isoform of PrP, PrPSc: Abnormal isoform of PrP.

related to the infection process.39,40 Since the ultimate detection limit should be the PrPC aggregate within a given sample volume, Bieschke et al. established a diagnostic test capable of detecting single aggregates of PrPSc.41 It was estimated that in the UK during the years from 1985 to 1995, tissue from more than 700,000 cattle carrying BSE entered the human food chain, while at the same time BSE was identified in 180,000 carcasses that were destroyed.42 This shows the necessity for sensitive methods that are able to identify BSE at an early stage. A sensitive paraffinembedded tissue (PET) blot method that detects prion PrPSc deposits in formalin-fixed and PET after blotting on a nitrocellulose membrane has been developed.43 Its sensitivity has

been compared with that of the histological and immunohistochemical (IHC) methods and the western blot method used in Prionics. With the PET blot method, four clinically less conspicuous cases showed the same PrPSc deposition pattern as was seen in clinical BSE. In these cases, the results of histological examination and western blotting were negative.43 Western blotting shows the presence and absence of protein production and also provides information on the molecular weight of peptides from the mobility shift of bands. Originally the application of a western blot technique for screening large numbers of cattle was considered difficult. Oesch et al. developed a rapid western blotting procedure (named Prionics Check or PWB), which has undergone

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­ odifications by the Swiss and British Veterinary Authorities m as well as the European Commission. Samples of brain stem from officially confirmed BSE or scrapie cases and samples from healthy New Zealand adult bovines could be diagnosed with 100% specificity and sensitivity.44,45 Furthermore, autolyzed samples could still be diagnosed correctly leading to the development of a unique surveillance program in Switzerland to test fallen stock for hidden cases of BSE. The routine application of the Prionics western blotting technique has also revealed unrecognized BSE cases that would have gone into the food chain. When peptides are modified with aberrant glycosylation, a different electrophoretic mobility is observed. Therefore, the mobility is influenced by the host genotype and also by prion strains.46,47 In other words, a mobility index is used to differentiate sCJD subtype (i.e., type 1 or type 2A), or iatrogenic CJD (iCJD) and sCJD to vCJD because of the different PrPSc glycoform profile.48–53 Recently, a modified western blotting method has been developed for prion detection. Cell blot54 is a method whereby cells grown on cover-slips are transferred to a nitrocellulose membrane where the PK-resistant PrP is then detected by western blotting.54 In addition, slot blotting55 is a method whereby cell lysates are filtered through a nitrocellulose membrane using a slot-blot device before the PK-resistant PrP in the membrane is detected by western blotting.55 The sensitivity of western blotting can be also enhanced by centrifugation56 or the development of an extraction method. An example of a modified extraction method is the sodium phosphotungstic acid (PTA) method.57,58 PTA can precipitate PrPSc from solutions. Guanidine hydrochloride has also been used to extract PrPSc. Western blotting has disadvantages in terms of processing time, the limited number of samples that can be processed, and the need for experienced personnel. These problems are at least in part overcome by the ELISA, although ELISA also has disadvantages (e.g., false-positive results and low cost-effectiveness), which western blotting does not have. Meanwhile, Grassi et al. chose to develop a conventional two-site immunometric assay (sandwich immunoassay) based on the use of two different monoclonal antibodies recognizing two distinct epitopes on the PrP molecule. In this category of immunoassay, the first antibody (capture antibody) is immobilized on a solid phase (e.g., a microtiter plate) while a second antibody, covalently labeled with an enzyme (e.g., acetylcholine esterase, AchE), is used as a tracer.59,60 PrP in a sample is detected by measuring enzyme activity bound to the solid phase through the intermediary of capture antibodyPrP-tracer antibody reactions. A two-site immunometric assay was chosen because it has been clearly demonstrated that this kind of immunoassay provides more sensitive, more specific, and faster measurement than other tests.61 With the ELISA method, PK-resistant PrP is directly absorbed on a microtiter plate or captured by the anti-PrP antibody coated on a microtiter plate, and detected or sandwiched by other anti-PrP antibodies. This mechanism of action is commercially exploited in Prionics Check EIA (Prionnics/Roche, Switzerland), Roche Applied Science PrionScreen (Roche, Switzerland), the Bio-Rad Platelia BSE purification kit and

Molecular Detection of Foodborne Pathogens

Bio-Rad BSE detection kit (Bio-Rad, France), the FRELISA BSE Kit (FUJIREBIO inc., Japan), the IDEXX HerdChek BSE antigen test kit (IDEXX Laboratories, USA), and Institut Pourquier Speed’it BSE (Institut Pourquier, France). Modified versions of this method are also commercially available such as VMRD CWD dbELISA (VMRD, Inc., USA), Enfer-TSE kit (Abbott Laboratories, USA), Prionics-Check LIA (Prionics, Switzerland), and CediTect BSE test (Eurofins Analytico Food, Netherlands). Although the extensively employed ELISA method is sensitive with high throughput and does not require sophisticated techniques, the high frequency of falsepositive results remains a problem. Another common method to diagnose prion diseases is the IHC analysis of brain sections (Figure 9.2).62,63 In IHC analysis, typical features of prion diseases such as the accumulation of PrP amyloid plaques, astroglyosis, and neuronal

BSE DMV F99-IHC 20x

BSE DMV 6C4-IHC 20x

BSE DMV T2-IHC 20x

Scrapie DMV F99 – IHC 20x

Scrapie DMV 6C4 – IHC 20x

Scrapie DMV T2 - IHC 20x

Figure 9.2  Immunohistochemistry of brain tissues from Italian cases of sheep scrapie and BSE. Diffuse particulate deposition of PrPSc was portrayed with mAbs 6C4 and T2 in the neuropil. The commercial mAb (F99/97) used as a control produced analogous results.(From Hosokawa, T. et al., Biochem. Biophys. Res. Commun. 366, 657, 2008. With permission from Dr. Casalone.)

Slow Viral Diseases

cell loss are examined by light microscopy. Although vacuolation is also sometimes used as an index of prion infection, many combinations of prion strains and host species portray PrPSc accumulation without vacuolation in brain sections after prion infection.64,65 Furthermore, the region of the brain where PrPSc is accumulated is dependent on the prion strain.66 Recently, the histopathological analysis of organs/ tissues other than the brain has been studied for the diagnosis of prion diseases. For example, the tonsils seem to be applicable in humans, deer, and sheep,58,67–72 while the appendix has been used for the preclinical diagnosis of vCJD in humans.58,73–76 Lesion profiles are often used for the discrimination of prion strains, which induces strain-specific patterns of vacuolation and PrPSc accumulation in the brain. In addition, IHC analysis has been improved with modified histoblotting or PET blotting. Such improvements further enable the detection of PrPSc in cryosections.43,77,78 There are several newly developed methods using antiPrP antibody for the detection of PrP or PrP isoforms. Conformation-dependent immunoassay (CDI) is a method employed to detect conformational differences between PrP isoforms by measuring the relative binding of antibodies to denatured and native proteins.57 CDI is commercially used for the CDI test in InPro Biotechnology, Inc. (USA). In other cases, using anti-PrP antibody-coupled microbeads and a flowcytometer, the flow microbead-immunoassay (FMI) method, detected 7-pmol/7-nmol recombinant PrP and PrPSc spiked in bovine meat and bone meal at concentrations higher than 0.3%.79 It should be emphasized that the novel PMCA technology amplifies cyclically the misfolding and aggregation process in vitro.80 It is conceptually analogous to DNA amplification by polymerase chain reaction (PCR). In this system, sequential cycles of incubation and sonication in the presence of seed PrPSc supplied with PrPC. Importantly, the PMCA method amplifies not only PrPSc but also prion infectivity titers.81 Furthermore, this method enables prion detection in blood from not only terminally diseased hamsters but also prion-infected presymptomatic hamsters.82 Five cycles of PMCA reaction achieved 97% conversion of PrPC to PrPSc in brain homogenate.83 This sensitivity is the highest among the detection methods for prion proteins reported so far. PrPSc detection is possible at >10,000-fold dilution, though the sensitivity could be increased if a more highly sensitive modified western blotting method was to be combined with this method. However, it should be noted that the protocol for PrPSc amplification depends on prion strain, species, and sample source (e.g., brain homogenate, blood, etc.). At present, the application of this method for vCJD, CWD, and hamster and mouse scrapie has been reported.83–86 Recently, the development of modified PMCA such as recombinant PrP-PMCA (rPrP-PMCA) has been studied.87 These developments will help to expand the use of this method in the field of prion biology. To date, this method has been mainly restricted to the study of prion conversion mechanisms and diagnosis of prion diseases. Capillary gel-electrophoresis is an approach that takes advantage of competitive antibody-binding between a

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fluorescein labeled synthetic PrP peptide and PrP present in tissue samples.88,89 The free peptide and antibody-peptide peaks are separated by capillary electrophoresis. The sensitivity of this method is high; only 50 amol/l (10 –18 mol/l) of fluorescent marker was used,89 while the specificity of this method remains unclear. For example, a recent report from Brown et al.90 has shown the difficulty in reproducing the results presented in a previous report by Schmerr et al.89 Fluorescence correlation spectroscopy (FCS) is also a highly sensitive method that detects single fluorescently labeled molecules in solutions.49,91 PrPSc can be labeled by the anti-PrP antibody or by conjugation with a labeled recombinant PrP. Fourier transformed infrared spectroscopy is a diagnostic method that incorporates the multivariate analysis of infrared spectra discriminating between prion-infected and uninfected animals.92 Such a spectroscopy-based assay is also used in multi-spectral ultraviolet fluorescence spectroscopy (MUFS).93 MUFS identifies changes in the spectral pattern of emission excited by ultraviolet radiation. The advantages of these methods are lack of pretreatment steps to eliminate PrPC (such as PK treatment) and anti-PrP antibody. However, the practical use of MUFS has not been reported so far. As similar spectral analysis, visible, and near-infrared (Vis-NIR) spectroscopy could be promising for noninvasive diagnosis94 because VisNIR radiation is transmissible into the body. Overall, discriminatory analysis of spectra between infected and uninfected animals could be useful for the diagnosis of prion diseases. A tool for detecting PrPSc has been developed recently. Although some antibodies and aptamers are able to distinguish PrPSc from PrPC, they have not had extensive practical applications. Antibodies with a conformational epitope of PrPSc were obtained by immunization of Tyr-Tyr-Arg peptide.95 Antibodies 15B3 and V5B2 can also recognize PrPScspecific epitopes.96,97 However, these antibodies have not been practically exploited and their applications in experiments have been limited. PrPC and/or PrPSc-specific binding RNA or DNA aptamers have also been reported.98–100 Although the use of animal bioassays has been considered the most sensitive way to detect infectivity, a recently developed western blot test is said to match or surpass the bioassays in reliability/accuracy.58 Although PrP-expressing transgenic mice inoculated with fibrils consisting of recombinant PrP or PMCA-amplified PrPSc exhibit a similar neuropathology similar to that found in prion diseases,2 it remains to be established whether PrPSc is completely identical in these cases or is the only entity of prions. Therefore, the determination of infectivity with a reliable animal bioassay is the gold standard, although the assay may require a longer time and many more animals to confirm this finding. Furthermore, the characteristics of this assay may occasionally demand sophisticated techniques.1 It should be also noted that the volume of inoculum is critical for reducing standard errors of the results in animal bioassays. Certain animal bioassays for prion diagnosis are time-consuming and economically nonviable. However, transmission of prions to their natural host is very informative and helpful for understanding the ­clinical phenotype of diseases. However, cell culture systems that

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specifically and reliably detect prion have been developed to certain strains of prion.101 Prion diseases can usually be diagnosed only by a postmortem analysis with an ELISA, western blotting, IHC means, and animal bioassays using brain samples.102 A premortem diagnostic method for prion diseases remains to be established. Recently, dramatic improvements to highly sensitive diagnostic methods such as PMCA have been achieved, some of which raise the possibility of a premortem diagnosis using blood samples, though the further development of analytical methods to specifically and reliably detect prions is more important with increasing emphasis on reducing prionrelated risks recognized in terms of public health and the safety of food and blood supplies. We still have the old questions for currently available immunological methods: (i) Which animal and in vitro models are currently available for the detection and quantification of prion infectivity? (ii) Which diagnostic techniques allow the reliable investigation and confirmation of suspect cases in animals and humans?

9.2  Methods 9.2.1 Reagents and Equipment Reagents: Dithiothreitol (DTT, Promega, Madison, WI) N-tetradecyl 1-N, N-dimethyl-3-ammonio-1-propanesulfonate, sulfobetaine, (SB 3-14, Calbiochem or Sigma) N-lauryl-sacrosine (Sigma), protease inhibitors were from Roche RNAse A (from bovine pancreas, protease-free) and DNAse I were from Calbiochem or Roche Protein concentration assay kit (BCA) from Pierce Others listed in Table 9.6. Equipment: To avoid cross contamination between different scrapie strains, we recommend the use of new glassware and instruments or single use materials for animal inoculations, dissection of brains from affected animals, and during the purification of PrPSc. This is especially important if the isolates are to be used in animal bioassays. Avoid blood contamination during dissection of brains, rinse excised brains in PBS, flash freeze in liquid nitrogen, and store at –70°C. All solutions must be filtered (0.2 μm) and placed in lint-free vessels previously rinsed with distilled or deionized water. All procedures are performed in a laminar flow biosafety cabinet in an appropriate laboratory according to governmental regulations. Use personal protection equipment (lab coat, gloves, and face mask) and change gloves frequently.

9.2.2 Sample Collection and Preparation PrPSc was prepared using a modified version of the procedure described by Shinagawa et al.103 PrPSc from TSE-affected brain tissue has been successfully purified in several species

Molecular Detection of Foodborne Pathogens

including humans with CJD and vCJD, sheep with scrapie, cattle with BSE, and cervids with CWD. Numerous experiments have used PrPSc isolates purified by this procedure allowing detailed analyses of several different aspects of TSE diseases. Typically the PrPSc yield from the hamster 263K strain is 50–100 μg per gram of brain. The method described yields PrPSc preparations with a total protein content of between 60 and 90%. Non-proteinase K (PK) digested preparations are usually less pure than PK-digested preparations since nearly all other proteins are digested by PK. However, a significant protein contaminant identified in these preparations is ferritin, which survives even PK treatment.104 Variations in purity may depend on the TSE strain, the stage of the disease, the manner in which the brain is excised and stored, and the expediency of the purification. In addition to contamination by other proteins, the final PrPSc sample most likely contains trace quantities of lipids, nucleic acids, carbohydrates, and ions (copper, iron, calcium, etc). Preparations not treated with PK are characterized by the presence of truncated and nontruncated PrPSc forms (Figure 9.3). This is likely due to in situ proteolysis that occurs in infected brains; however, additional proteolysis may occur during the excision and/or purification procedure even though steps such as rapid freezing, careful temperature control, and the use of protease inhibitors during the purification are used in an attempt to prevent proteolysis. Since PrPSc itself is thought to be the infectious agent of TSEs, it would be of interest to achieve its purification to homogeneity in order to test this hypothesis.

9.2.3  Detection Procedures PrPSc is the most reliable biomarker for prion diseases and also the main component of prions. Therefore, most of the methods used for the diagnosis of prion infections rely on the presence of PrPSc (Table 9.5). Following PK digestion PrPC is completely degraded, whereas the C-terminal of fragment PrPSc is partially resistant. This feature is used to distinguish PrPSc from PrPC.37,105 As PrPSc is highly accumulated in brain in the clinical phase of diseases, brain samples are appropriate for these methods, which are required for post-mortem analysis. One of the most reliable methods of detecting prions is western blotting. Western blotting is used to detect PK-resistant PrP in extracts from the brain and other tissues. PK-digested samples are first subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), followed by transfer of the separated peptides blotted onto the membrane with the anti-PrP antibody. A western blotting kit (Prionics Check Western Test) is commercially available from Prionics AG (Switzerland). To cite a case, a representative simple step-wise protocol of western blotting for prion-noninfected HpL3-4-PrP cells106 and persistently prion-infected ScN2a cells with anti-PrP antibody SAF83 (SPI Bio., Montigny le Bretonneux, France) has been shown as follows:

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Slow Viral Diseases

Table 9.6 Solutions, Reagents, and Buffers Buffer

Component

Volume

10 × PBS pH 6.9

26.8 g Na2HPO4-7H2O 14.8 g NaH2PO4-H2O 75.9 g NaCl

1000 ml

1 × PBS, pH 7.4

100 ml 10× PBS, pH 6.9 900 ml de-ionized water 340.5 ml M Na2HPO4 159.5 ml M NaH2PO4 10 ml 1 M Tris-Cl, pH 8.0 2 ml 0.5 M EDTA 26.2 ml 5 M NaCl 0.002 ml 1 M DTT 50 g Na N-lauryl-sacrosine 100 ml 5× TEND 171 ml 5 M NaCl 100 ml 5× TEND 5 g SB 3–14 1 ml 1 M Tris-Cl 0.5 ml 1 M MgCl2 2 ml 5 M NaCl 68.5 g sucrose 4 ml 5 M NaCl 1 g SB 3–14 2 ml 1 M Tris-Cl 0.5 g SB 3–14 10 ml 10× PBS

1000 ml

1 M NaPO4, pH 6.9 5 × TEND (50 mM Tris-HCl, pH 8.0 at 4°C; 5 mM EDTA; 5 mM EDTA; 665 mM NaCl; 1 mM DTT, added immediately before use) 10% Sarcosine (TEND) 10% NaCl, 1% SB 3-14 (TEND)

TMS (10 mM Tris-HCl, pH 7.0 at 20°C; 10 mM Tris-HCl, pH 7.0 at 20°C; 100 mM NaCl) 1 M Sucrose; 100 mM NaCl; 0.5% SB 3–14; 10 mM Tris-CI, pH 7.4 at 20°C

0.5% SB 3–14/1× PBS

500 ml 500 ml

500 ml

100 ml

200 ml

100 ml

1 M DTT (1000 ×) Protease inhibitors Pls (1000 ×)

0.5 mg/ml Leupeptin = 1 μM (in H2O) 1 mg/ml Aprotinin = 0.15 μM (in H2O) 1 mg/ml Pepstatin = 1 μM (in methanol) 24 mg/ml Pefabloc = 0.1 M (in 0.05 M Tris-HCl, pH 6.5) at 20°C

Note: DTT and PI solutions are freshly diluted into buffers.

(1) For preparation of the cell lysate, detach cells with a scraper and wash twice with ice-cold PBS. (2) Solubilize the washed cells in radio-immunoprecipitation assay (RIPA) buffer containing 10 mM TrisHCl (pH 7.4), 1% deoxycholate, 1% Nonidet P-40, 0.1% sodium dodecyl sulfate (SDS), and 150 mM NaCl and then sonicated at 4°C for 10 min. (3) Remove the cellular debris by centrifugation at 5,000 × g for 1 min. (4) Measure the protein concentration of the supernatants using a Bio-Rad DC protein assay (Bio-Rad, Hercules, CA). (5) Treat the sample (120 μg protein) with proteinase K at 20 μg/ml for 30 min at 37°C. (6) Add an equal volume of 2× SDS gel-loading buffer (90 mM Tris/HCl pH 6.8, 10% mercaptoethanol, 2% SDS, 0.02% bromophenol blue, and 20% glycerol) and heat the samples at 100°C for 10 min to terminate the reaction before western blotting. Cells treated as above except for the digestion by PK should be also included as control.

(7) Separate the proteins by conventional SDS-PAGE (12%) as described elsewhere. (8) Transfer the proteins to polyvinylidene difluoride (PVDF) membranes (Amersham Biosciences, Piscataway, NJ) using a semidry blotting system (Bio Rad, Cambridge, MA). (9) Block the membranes with 5% skim milk for 1 h at room temperature. (10) Further incubate the blocked membranes for 1 h at room temperature with anti-PrP antibody SAF83 (SPI bio), which recognizes residues 126–164 of PrP, in PBS-Tween (0.1% Tween 20) containing 0.5% skim milk. Note: anti-PrP antibodies recognizing the C-terminal half of PrP such as SAF83 and 6H4 (Prionics AG) can be used for the detection of PrPSc in this western blotting system, whereas those recognizing the N-terminal half of PrP such as SAF32 (SPI bio) cannot be used. This is because the C-terminal half of PrPSc is resistant to PK, whereas the N-terminal half is susceptible to PK.

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Molecular Detection of Foodborne Pathogens

1

2

3

Uninfected PK

–

+

4

Infected –

+

5

6

7

Uninfected –

8

Infected

+

–

+

HpL3-4-PrP (Prion non-infected cells) (kDa) 51

PK

(–)

ScN2a (Persistently prion infected cells)

(+)

PK

(–)

(+)

75 50 37

35 29

25 20

Anti-PrP Ab (SAF83)

15

10 21 6H4

P8

Figure 9.3  Detection of PrPSc in the brains of terminally diseased mice. PrP from the brain membrane fraction (30 μg protein per lane) of uninfected mice (lanes 1, 2, 5, 6) and terminally diseased mice intracerebrally inoculated with Obihiro-1 prions (lanes 3, 4, 7, 8) was detected by western blotting with anti-PrP 6H4 (lanes 1–4) and P8 (lanes 5–8). Each fraction was treated (+)(lanes 2, 4, 6, 8) with proteinase K (PK), or left untreated (–) (lanes 1, 3, 5, 7). (From Inoue, Y. et al., Jpn J. Infect. Dis., 58, 78, 2005. With permission from the National Institute of Infectious Diseases, Japan.)

(1) Wash the membranes three times for 10 min in PBSTween and incubated with horseradish peroxidase (HRP)-labeled anti-mouse immunoglobulin secondary antibody in PBS-Tween containing 0.5% skim milk for 1 h at room temperature before being washed three times in PBS-Tween for 10 min. (2) Further develop the membranes with an enhanced chemiluminescence (ECL) reagent (Amersham) for 5 min. Blots were exposed to ECL Hypermax film (Amersham). Films were processed in an X-ray film processor. The results obtained by western blotting using HpL3-4-PrP and ScN2a cells with SAF83 are shown in Figure 9.4. HpL34-PrP cells106 generate PrPC but not PrPSc, while ScN2a cells produce both PrPC and PrPSc. Therefore, PrP in HpL3-4-PrP cells is digestible with PK, whereas PrP in ScN2a is partially resistant to PK. Therefore, it is obvious that western blotting of PrP using the anti-PrP antibody detects PrP in ScN2a but not in HpL3-4-PrP after digestion by PK (Figure 9.4).

9.3 Conclusions and Future Perspectives The chain reaction of BSE epidemics in the UK and Europe and subsequent emergence of vCJD in young adults and teenagers have raised concerns and highlighted the importance of risk assessment in the food chain. Recently, several highly

Figure 9.4  Detection of the abnormal isoform of prion protein (PrPSc) in cell cultures. Detection of prion protein (PrP) in lysates from HpL3-4 cells (PrP gene-deficient neuronal cells retransformed with a PrP-expressing vector) and ScN2a cells (persistently prion-infected neuroblastoma) with (+) and without (–) proteinase K (PK) treatment was performed by western blotting with anti-PrP antibody SAF83 (SPI Bio, Montigny le Bretonneux, France). PK digested the cellular isoform of prion protein (PrPC) without affecting PrPSc. Therefore, PK-resistant PrP is usually used as an index of prion infection. A relevant example of PK-resistant PrP detected in ScN2a cells but not in HpL3-4-PrP cells is shown.

sensitive methods for detecting prions have been developed. Representative of these is PMCA. Originally developed by Claudio Soto and his colleagues, PMCA has been a hot topic of debate in prion meetings all over the world. A broad spectrum of PrPSc species have now been successfully amplified using PMCA, including CWD, mouse-adapted scrapie, and BSE. Studies with human sporadic and variant CJD cases show that PMCA amplification efficiency is tightly controlled by the PrPC substrate genotype at codon 129. PMCA appears to overcome the species barrier encountered during cross-species transmission more rapidly than in vivo. By “forcing” the technique with lower dilutions of the PrPSc seed and more amplification rounds, mouse Chandler PrPSc can now convert hamster PrPC or cervid PrPC, a conversion that might be observable in vivo but only with extremely long incubation periods. Castilla J. showed that using PMCA, PrPSc was generated from the healthy brains of 11 different species, including bank voles, mice, cattle, humans, sheep, and rabbits, generating a variety of electrophoretic profiles.107 PMCA was able to detect PrPSc in as little as 1 μl of blood from an asymptomatic prion-infected mouse. Given the increasing evidence of human to human transmission via blood products, scientists are waiting for PMCA to be incorporated into a reliable test with the ability to identify blood donors that are asymptomatic carriers. Further advances in amplification technology are to be expected and the replacement of PrPC by recombinant PrP as a substrate as well as the use of intermittent shaking rather than sonication should circumvent some of the difficulties in the near future.

Slow Viral Diseases

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124 48. Collinge, J. et al. Molecular analysis of prion strain variation and the aetiology of ‘new variant’ CJD. Nature, 383, 685, 1996. 49. Head, M.W. et al. Prion protein heterogeneity in sporadic but not variant Creutzfeldt-Jakob disease: UK cases 1991–2002. Ann. Neurol., 55, 851, 2004. 50. Notari, S. et al. Effects of different experimental conditions on the PrPSc core generated by protease digestion: implications for strain typing and molecular classification of CJD. J. Biol. Chem., 279, 16797, 2004. 51. Parchi, P. et al. Classification of sporadic Creutzfeldt-Jakob disease based on molecular and phenotypic analysis of 300 subjects. Ann. Neurol., 46, 224, 1999. 52. Schoch, G. et al. Analysis of prion strains by PrPSc profiling in sporadic Creutzfeldt-Jakob disease. PLoS Med., 3, e14, 2006. 53. Wadsworth, J.D. et al. Molecular and clinical classification of human prion disease. Br. Med. Bull., 66, 241, 2003. 54. Bosque, P.J. and Prusiner, S.B. Cultured cell sublines highly susceptible to prion infection. J. Virol., 74, 4377, 2000. 55. Winklhofer, K.F., Hartl, F.U. and Tatzelt, J. A sensitive filter retention assay for the detection of PrP(Sc) and the screening of anti-prion compounds. FEBS Lett., 503, 41, 2001. 56. Fujita, R. et al. Efficient detection of PrPSc (263K) in human plasma. Biologicals, 34, 187, 2006. 57. Safar, J. et al. Eight prion strains have PrP(Sc) molecules with different conformations. Nat. Med., 4, 1157, 1998. 58. Wadsworth, J.D. et al. Tissue distribution of protease resistant prion protein in variant Creutzfeldt-Jakob disease using a highly sensitive immunoblotting assay. Lancet, 358, 171, 2001. 59. Ekins, R.P. and Jackson, T. Non isotopic immunoassays. In: Monoclonal Antibodies and New Trends in Immunoassay. An Overview, C.A. Bizollon (Ed.). p. 129. Elsevier, Amsterdam, 1984. 60. Grassi, J. et al. Production of monoclonal antibodies against interleukin-1 alpha and -1 beta. Development of two enzyme immunometric assays (EIA) using acetylcholinesterase and their application to biological media. J. Immunol. Methods, 123, 193, 1989. 61. Moynagh, J. et al. Tests for BSE evaluated. Bovine spongiform encephalopathy. Nature, 400, 105, 1999. 62. Armstrong, R.A. et al. Quantification of vacuolation (“spongiform change”), surviving neurones and prion protein deposition in eleven cases of variant Creutzfeldt-Jakob disease. Neuropathol. Appl. Neurobiol., 28, 129, 2002. 63. Ironside, J.W. et al. Pathological diagnosis of variant Creutzfeldt-Jakob disease. APMIS, 110, 79, 2002. 64. Iwata, N. et al. Distribution of PrP(Sc) in cattle with bovine spongiform encephalopathy slaughtered at abattoirs in Japan. Jpn J. Infect. Dis., 59, 100, 2006. 65. Orge, L. et al. Identification of putative atypical scrapie in sheep in Portugal. J. Gen. Virol., 85, 3487, 2004. 66. Hirogari, Y. et al. Two different scrapie prions isolated in Japanese sheep flocks. Microbiol. Immunol., 47, 871, 2003. 67. Hill, A.F. et al. Diagnosis of new variant Creutzfeldt-Jakob disease by tonsil biopsy. Lancet, 349, 99, 1997. 68. Hill, A.F. et al. Investigation of variant Creutzfeldt-Jakob disease and other human prion diseases with tonsil biopsy samples. Lancet, 353, 183, 1999. 69. Schreuder, B.E. and Somerville, R.A. Bovine spongiform encephalopathy in sheep? Rev. Sci. Tech., 22, 103, 2003. 70. Schreuder, B.E. et al. Preclinical test for prion diseases. Nature, 381, 563, 1996. 71. van Keulen, L.J. et al. Pathogenesis of natural scrapie in sheep. Arch. Virol. Suppl., 57, 2000.

Molecular Detection of Foodborne Pathogens 72. Wild, M.A. et al. Preclinical diagnosis of chronic wasting ­disease in captive mule deer (Odocoileus hemionus) and white-tailed deer (Odocoileus virginianus) using tonsillar biopsy. J. Gen. Virol., 83, 2629, 2002. 73. Hilton, D.A. et al. Prion immunoreactivity in appendix before clinical onset of variant Creutzfeldt-Jakob disease Lancet, 352, 703, 1998. 74. Hilton, D.A. et al. Accumulation of prion protein in tonsil and appendix: review of tissue samples. BMJ, 325, 633, 2002. 75. Hilton, D.A. et al. Prevalence of lymphoreticular prion protein accumulation in UK tissue samples. J. Pathol., 203, 733, 2004. 76. Ironside, J.W. et al. Retrospective study of prion-protein accumulation in tonsil and appendix tissues. Lancet, 355, 1693, 2000. 77. Ritchie, D.L., Head, M.W. and Ironside, J.W. Advances in the detection of prion protein in peripheral tissues of variant Creutzfeldt-Jakob disease patients using paraffin-embedded tissue blotting. Neuropathol. Appl. Neurobiol., 30, 360, 2004. 78. Taraboulos, A. et al. Regional mapping of prion proteins in brain. Proc. Natl. Acad. Sci. USA, 89, 7620, 1992. 79. Murayama, Y. et al. Specific detection of prion antigenic determinants retained in bovine meat and bone meal by flow microbead immunoassay. J. Appl. Microbiol., 101, 369, 2006. 80. Soto, C., Saborio, G.P. and Anderes, L. Cyclic amplification of protein misfolding: application to prion-related disorders and beyond. Trends Neurosci., 25, 390, 2002. 81. Castilla, J. et al. In vitro generation of infectious scrapie prions. Cell, 121, 195, 2005. 82. Saa, P., Castilla J. and Soto, C. Presymptomatic detection of prions in blood. Science, 313, 92, 2006. 83. Saborio, G.P., Permanne, B. and Soto, C. Sensitive detection of pathological prion protein by cyclic amplification of protein misfolding. Nature, 411, 810, 2001. 84. Jones, M. et al. In vitro amplification and detection of variant Creutzfeldt-Jakob disease PrPSc. J. Pathol., 213, 21, 2007. 85. Kurt, T.D. et al. Efficient in vitro amplification of chronic wasting disease PrPRES. J. Virol., 81, 9605, 2007. 86. Murayama, Y. et al. Efficient in vitro amplification of a mouse-adapted scrapie prion protein. Neurosci. Lett., 413, 270, 2007. 87. Atarashi, R. et al. Ultrasensitive detection of scrapie prion protein using seeded conversion of recombinant prion protein. Nat. Methods, 4, 645, 2007. 88. Jackman, R. and Schmerr, M.J. Analysis of the performance of antibody capture methods using fluorescent peptides with capillary zone electrophoresis with laser-induced fluorescence. Electrophoresis, 24, 892, 2003. 89. Schmerr, M.J. et al. Use of capillary electrophoresis and fluorescent labeled peptides to detect the abnormal prion protein in the blood of animals that are infected with a transmissible spongiform encephalopathy. J. Chromatogr. A, 853, 207, 1999. 90. Brown, P., Cervenakova, L. and Diringer, H. Blood infectivity and the prospects for a diagnostic screening test in CreutzfeldtJakob disease. J. Lab. Clin. Med., 137, 5, 2001. 91. Giese, A. et al. Putting prions into focus: application of single molecule detection to the diagnosis of prion diseases. Arch. Virol. Suppl., 161, 2000. 92. Schmitt, J. et al. Identification of scrapie infection from blood serum by Fourier transform infrared spectroscopy. Anal. Chem., 74, 3865, 2002.

Slow Viral Diseases 93. Rubenstein, R. et al. Detection and discrimination of PrPSc by multi-spectral ultraviolet fluorescence. Biochem. Biophys. Res. Commun., 246, 100, 1998. 94. Sakudo, A. et al. Near-infrared spectroscopy: promising diagnostic tool for viral infections. Biochem. Biophys. Res. Commun., 341, 279, 2006. 95. Paramithiotis, E. et al. A prion protein epitope selective for the pathologically misfolded conformation. Nat. Med., 9, 893, 2003. 96. Curin Serbec, V. et al. Monoclonal antibody against a peptide of human prion protein discriminates between CreutzfeldtJacob’s disease-affected and normal brain tissue. J. Biol. Chem., 279, 3694, 2004. 97. Korth, C. et al. Prion (PrPSc)-specific epitope defined by a monoclonal antibody. Nature, 390, 74, 1997. 98. Rhie, A. et al. Characterization of 2’-fluoro-RNA aptamers that bind preferentially to disease-associated conformations of prion protein and inhibit conversion. J. Biol. Chem., 278, 39697, 2003. 99. Sayer, N.M. et al. Structural determinants of conformationally selective, prion-binding aptamers. J. Biol. Chem., 279, 13102, 2004. 100. Weiss, S. et al. RNA aptamers specifically interact with the prion protein PrP. J. Virol., 71, 8790, 1997. 101. Sakudo, A. et al. Recent developments in prion disease research: diagnostic tools and in vitro cell culture models. J. Vet. Med. Sci., 69, 329, 2007. 102. Gavier-Widen, D. et al. Diagnosis of transmissible spongiform encephalopathies in animals: a review. J. Vet. Diagn. Invest., 17, 509, 2005. 103. Onodera, T. et al. Isolation of scrapie agent from the placenta of sheep with natural scrapie in Japan. Microbiol. Immunol., 37, 311, 1993. 104. Bolton, D.C. et al. Isolation and structural studies of the intact scrapie agent protein. Arch. Biochem. Biophys., 258, 579, 1987. 105. McKinley, M.P., Bolton, D.C. and Prusiner, S.B. A proteaseresistant protein is a structural component of the scrapie prion. Cell, 35, 57, 1983. 106. Sakudo, A. et al. Octapeptide repeat region and N-terminal half of hydrophobic region of prion protein (PrP) mediate PrP-dependent activation of superoxide dismutase. Biochem. Biophys. Res. Commun., 326, 600, 2005.

125 107. Castilla, J. et al. De novo generation of prions in a cell-free system. Presented at Prion2007, Edinburgh, September 26–28, 2007, 16. 108. Inoue, Y. et al. Infection route-independent accumulation of splenic abnormal prion protein. Jpn J. Infect. Dis., 58, 78, 2005. 109. Grathwohl, K.U. et al. Sensitive enzyme-linked immunosorbent assay for detection of PrP(Sc) in crude tissue extracts from scrapie-affected mice. J. Virol. Methods, 64, 205, 1997. 110. McBride, P.A., Bruce, M.E. and Fraser, H. Immunostaining of scrapie cerebral amyloid plaques with antisera raised to scrapie-associated fibrils (SAF). Neuropathol. Appl. Neurobiol., 14, 325, 1988. 111. Prusiner, S.B. et al. Measurement of the scrapie agent using an incubation time interval assay. Ann. Neurol., 11, 353, 1982. 112. Klohn, P.C. et al. A quantitative, highly sensitive cell-based infectivity assay for mouse scrapie prions. Proc. Natl. Acad. Sci. USA, 100, 11666, 2003. 113. Barnard, G. et al. The measurement of prion protein in bovine brain tissue using differential extraction and DELFIA as a diagnostic test for BSE. Luminescence, 15, 357, 2000. 114. Fischer, M.B. et al. Binding of disease-associated prion protein to plasminogen. Nature, 408, 479, 2000. 115. Kenney, K. et al. An enzyme-linked immunosorbent assay to quantify 14-3-3 proteins in the cerebrospinal fluid of suspected Creutzfeldt-Jakob disease patients. Ann. Neurol., 48, 395, 2000. 116. Miele, G., Manson, J. and Clinton, M. A novel erythroidspecific marker of transmissible spongiform encephalopathies. Nat. Med., 7, 361, 2001. 117. Zerr, I. et al. Diagnosis of Creutzfeldt-Jakob disease by twodimensional gel electrophoresis of cerebrospinal fluid. Lancet, 348, 846, 1996. 118. Hosokawa, T., et al. Distinct immunohistochemical localization of Kuru plaques using novel anti-PrP antibodies. Microbiol. Immunol., 52, 25, 2008. 119. Hosokawa, T. et al. A monoclonal antibody (1D12) defines novel distribution pattern of prion protein (PrP) as granules in nucleus: implication for mechanisms of nuclear localization of PrP. Biochem. Biophys. Res. Commun. 366, 657, 2008.

Section II Foodborne Gram-Positive Bacteria

10 Bacillus

Noura Raddadi, Aurora Rizzi, Lorenzo Brusetti, Sara Borin, Isabella Tamagnini, and Daniele Daffonchio University of Milan

Contents 10.1 Introduction ................................................................................................................................................................... 129 10.1.1 Classification..................................................................................................................................................... 129 10.1.2 The Genetic Baseline of the B. cereus Group Ecotypes................................................................................... 130 10.1.3 Species Discrimination in the B. cereus Group................................................................................................ 130 10.1.4 Emetic and Diarrheal Illnesses Caused by Foodborne B. cereus......................................................................131 10.1.5 Other Foodborne Virulent Bacillus spp............................................................................................................ 133 10.1.6 Detection of B. cereus Toxins........................................................................................................................... 133 10.1.7 Detection of B. cereus Cells in Food and Environmental Samples.................................................................. 134 10.2 Methods.......................................................................................................................................................................... 135 10.2.1 Sample Preparation........................................................................................................................................... 135 10.2.1.1 B. cereus Isolation and Enumeration............................................................................................... 135 10.2.1.2 DNA Isolation.................................................................................................................................. 136 10.2.2 Detection Procedures........................................................................................................................................ 136 10.2.2.1 Detection of B. anthracis by RSR-PCR ......................................................................................... 136 10.2.2.2 Detection of B. cereus Toxin Genes by Multiplex PCR.................................................................. 137 10.2.2.3 Detection of B. cereus Toxin Genes by RT-PCR............................................................................. 138 10.2.2.4 Detection of Emetic B. cereus Strains by RT-PCR......................................................................... 138 10.3 Conclusions and Future Perspectives............................................................................................................................. 139 References.................................................................................................................................................................................. 140

10.1  Introduction 10.1.1 Classification The genus Bacillus includes Gram-positive spore-forming rod-shaped bacteria that are very diverse, from physiology to the ecological niche, from DNA sequence to gene regulation. Several species within the genus Bacillus have been associated with foodborne diseases, or carrying toxin determinants: among others, B. weihenstephanenesis, B. pumilus, B. mojavensis, B. licheniformis, and B. subtilis.1–5 However, the best-known foodborne pathogen within the genus is B. cereus. All Bacillus species, owing to endospore formation, can survive heat treatment and in the absence of a competitive microbiota, can multiply in cooked food during storage, producing toxins under suitable conditions.6–8 Consumption of such contaminated food might lead to intoxication with acute symptoms shortly after ingestion. Bacterial cells and/or spores in the food can survive the passage through the acidic environment of the stomach and infect the intestine with subsequent toxin production and symptoms, which last hours to days after food consumption.5 In this chapter, we will focus on the food-poisoning implications of Bacillus species. Before giving details on the food poisoning potential, the type of toxins and the method

of detection, we will briefly introduce the complex ecology and the identification challenges that characterize several Bacillus species. We will use as a paradigm the B. cereus group that shows a ‘bivalent face’ as far as its important impact on human activity.9,10 B. cereus is the most important foodborne pathogen in this group and will be the major subject of this chapter. This bacterium has been associated with food-poisoning illnesses11–13 as well as other kinds of clinical infections.14–18 B. thuringiensis is an insect pathogen widely used as biopesticide and differently from B. cereus has a very useful impact on human activities being widely used in agriculture for insect pest biocontrol. B. anthracis is the etiological agent of anthrax, a fatal disease for human and animals and has been sometimes associated with foodborne anthrax cases.19–21 Three forms of anthrax can be distinguished depending on the route of infection: cutaneous, gastrointestinal, and inhalational (pulmonary). Each form may progress to fatal systemic anthrax. The gastrointestinal form is extremely rare and occurs frequently after the ingestion of undercooked meat from animals with B. anthracis. The means by which B. anthracis crosses membrane barriers to establish infection remains unknown. B. anthracis probably invades the mucosa through 129

130

preexisting lesions, or, in the absence of mucosal damage, through the Peyer’s patch due to the uptake by M cells or dendritic cells. The disease is characterized by fever, nausea, vomiting, abdominal pain, and bloody diarrhea.19,22 B. weihenstephanensis, a psychrotolerant species capable of growing at temperatures as low as 4–6°C is implicated in food spoilage.23,24 In addition to B. cereus, B. thuringiensis, B. anthracis, and B. weihenstaephanenesis, the B. cereus group encompasses two other species, B. mycoides and B. pseudomycoides that are typically isolated from soil and plant rhizosphere. The six species are known to be strictly related phylogenetically, as has been shown by DNA-DNA hybridization studies23,25–27 and the sequencing of the ribosomal RNA genes.23,28–30 However, a marked variability is always observed when large collections of strains are examined by DNA fingerprinting methods that target the whole genome31–35 and/or discrete genes.36–40 Hence, the phylogenetic and taxonomic relationship among these species is still open to debate. It has been proposed previously that B. anthracis, B. cereus, and B. thuringiensis represent a single species, this conclusion having been reached through genome sizing and mapping,32–35 multilocus enzyme electrophoresis (MEE),41 multilocus sequence typing (MLST),42,43 and genome sequencing.44 The genetic variability observed within/between these species has raised several interesting questions and posed a challenge to microbiologists: (i) What is the evolutionary pathway of these species and how are they differentiated during evolution? (ii) What is the genetic baseline driving the different ecotypes and the virulence?

10.1.2 The Genetic Baseline of the B. cereus Group Ecotypes The ecology of these bacteria is still far to be clear, despite several analyses have recently shown that a major environmental niche for these bacteria is the invertebrate intestinal tract.45–47 In a general model, these species commonly live associated to the intestine of invertebrates assuming a symbiotic life style, but occasionally they escape such an ecotype becoming invasive for other animal hosts, that are in specific insects, arthropods, and nematodes for B. thur­ ingiensis, mammals, and humans for B. anthracis and B. cereus.46 Apart from this general scheme, the ecology of these species outside the invertebrate host is not completely clear, and several environments where these bacteria have been commonly associated to, have been questioned to be really able to continuously support the life cycle of these bacteria. It has been shown that with respect to other soil inhabiting bacteria, like those of the B. subtilis group that have genomes harboring many genetic determinants for the metabolism of plant-derived sugars, B. cereus group bacteria are rich in genes for protein metabolism. This suggests that they evolved in relation to nutrient rich environments like animal guts or animal tissues and fluids rather than the plant-environment.44,48 Specialization on different “animal” niches has lead to different evolution of the species in the

Molecular Detection of Foodborne Pathogens

B. cereus group. For instance, B. anthracis has been proposed to diverge from the other members of the group by specializing as a lethal mammal pathogen. It has been supposed that B. anthracis has evolved along two possible pathways:49 (i) in the first pathway, one considers B. anthracis to be a relatively ancient organism with a low growth rate, determined by the previously mentioned ecological constraints and evolving separately from a common ancestor with B. cereus and other relatives; (ii) in the second pathway, it has been supposed to be derived relatively recently from B. cereus, through the acquisition and rearrangement of plasmids resulting in the actual pXO plasmid pattern responsible of lethal virulence. Such a divergence from the B. cereus ancestor has been proposed to occur rather recently, between 13,000 and 26,000 years ago.50 A recent divergence from B. cereus is supported by (i) the very marked similarity between B. anthracis and certain strains of B. cereus;41,51 (ii) the presence in B. anthracis genome of several B. cereus-typical virulence genes that are, however, not expressed.52 The lack of gene expression in B. anthracis has been shown to be due to a nonsense mutation in PlcR, a pleiotrophic regulator driving the transcription of several B. cereus virulence factors (such as phospholipase C, proteases and enterotoxins) except cereulide.53–55 Recently, it has been shown that under anaerobiosis, at least two separate pathways, i.e., the PlcR-dependent and the Fnr-dependent pathways, control the expression of enterotoxin genes.56–58 The activity of PlcR depends on PapR, a secreted signaling peptide reimported into the bacterial cell through the oligopeptide permease (Opp) system59 while the activity of Fnr depends on the availability of O2 and NO. In B. anthracis, a nonfunctional PlcR is the result of counter selection, due to its disadvantageous effects on the overall fitness of the cell. Mignot et al.52 showed that a functional PlcR is incompatible with the plasmid borne AtxA-controlled virulence regulon, determining a dramatic effect on the sporulation. When the two regulators are simultaneously transcribed B. anthracis cells loose the ability to sporulate, a feature that heavily affects the overall survival capacity of the bacterium in the environment.

10.1.3 Species Discrimination in the B. cereus Group The similarity among such closely related bacteria as the species of the B. cereus group posed serious challenges for the species discrimination. This is an important point to be addressed, considering the differences in the virulence potential of these microorganisms. While B. anthracis can be differentiated by biochemical tests from most of B. cereus and B. thuringiensis, a problem still exists for isolates borderline between species such as, for instance, the pathogenic B. cereus G9241, for which these tests fail to recognize its pathogenic potential.60 This strain has been confirmed to be a B. cereus harboring a virulence plasmid very similar to plasmid pXO1 of B. anthracis, and a second plasmid encoding for a capsule synthesis. However, such a second plasmid and the capsule coding genes were completely different from the pXO2 plasmid and the typical capsule of B. anthracis.60

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Bacillus

Thus, considering such a virulence potential of strains genetically near neighbor of B. anthracis, approaches that might rapidly identify these strains are of great interest. Indeed the characterization of these strains is actually providing useful clues for the understanding of the evolution pattern of these species and the mechanisms they have adopted for regulating the virulence by the way of plasmid/chromosome interaction.52 From a safety perspective, these strains could represent alternative hosts for B. anthracis toxin genes.60 Several B. cereus strains resulted strictly related to B. anthracis. For example, Keim et al.49 and Radnedge et al.61 individuated by amplified fragment length polymorphism (AFLP), B. cereus and B. thuringiensis strains closely related to B. anthracis. Radnedge et al.61 tried to identify by suppression subtractive hybridization genomic regions of B. anthracis absent in these closely related strains. Apart from AFLP, several other methods based on whole genome fingerprinting have been used for typing B. anthracis and the identification of species borderline strains, like, among others, rep-PCR10 and multi locus sequence typing (MLST),42 or, recently, comparative genomics62 and microarray analysis.63 Alternative approaches have been based on length or sequence polymorphisms in variable number tandem repeats in multiple loci (multi locus VNTR analysis, MLVA64) or on signature single nucleotide polymorphisms (SNPs) in the genome.65 MLVA has been used as a gold standard for subtyping B. anthracis isolated worldwide (see among others Refs 64, 66–68). Together with MLVA, SNP analysis is greatly contributing in typing and tracing B. anthracis isolates, especially by the whole genome SNP analysis.50 Besides those identified in the whole genome, SNPs in housekeeping genes have been nicely exploited to identify strains related to certain species or genetic types. For example, Prüss et al.69 showed that certain nucleotides in the 16S rRNA gene and their relative prevalence among the different ribosomal operons in the genome correlate with the psychrotolerance of B. cereus strains and hence are signature of the species B. weihenstephanensis. In addition to strain B. cereus G9241, several other isolates have been shown to be borderline between B. cereus or B. thuringiensis and B. anthracis. In population genetic studies among a strain collection of the B. cereus group species it was found MEE and MLST41,42 that the strains could be grouped into two main groups, the first including soil and dairy isolates, while the second those with pathogenic potential. This second group included, besides B. anthracis, most of the strains isolated from patients in clinical environments and, among these strains, the B. cereus strains isolated from periodontitis in humans. Helgason et al.41,42 clearly showed that in the species B. cereus/B. thuringiensis some genetic types exist having a close relationship with B. anthracis. For instance, besides the previously mentioned strain G9241, B. cereus Zebra Killer (ZK) and B. thuringiensis 97–27 (subsp. Konkukian, serotype H34)70 resulted closely related to B. anthracis according to AFLP and MLST.71 The proteome of all these three isolates resulted more similar to that of B. anthracis than to the nonvirulent B. cereus ATCC 14579.72 In a study aiming to determine a strategy for the identification

of B. cereus group isolates near neighbor of B. anthracis,43 two B. cereus and one B. thuringiensis strains resulted closely related to B. anthracis, according to restriction site insertion (RSI)-PCR36 patterns of the 16S–23S rRNA gene spacers, rep-PCR profiles and MLST analysis.43 Despite several approaches based on comparative genome sequencing and SNPs analysis by microarray applications, have been recently proposed for the identification of strains close to B. anthracis and other virulent strains in the B. cereus group, the gold standard is actually represented by MLST. This approach is rather simple and straightforward and take advantage from the availability of databases for the comparison of new isolates with historical or well characterized strains.42,73 Several MLST schemes based on sequencing of different gene sets have been proposed.42,43,73–77 However, most of the published studies are referring to that developed by Priest et al.74 From this MLST scheme, B. anthracis fall within the clade 1 that also includes many virulent B. cereus near neighbors of B. anthracis including emetic strains.

10.1.4 Emetic and Diarrheal Illnesses Caused by Foodborne B. cereus This bacterium is a common soil inhabitant that has been recognized as a food poisoning species since the beginning of the century. The first confirmed outbreak of B. cereus food poisoning occurred in Norway in 1950 after the consumption of a contaminated vanilla sauce, which had been prepared a day in advance before consumption and stored at room temperature.78 B. cereus is widespread in the environment including soil, water, and phylloplane.79 It is also often present in a variety of foods including rice, cereals, spices, vegetables, meat, milk, and pasteurized milk products.80–87 Toxins produced by vegetative cells of B. cereus can be the causative agents of two types of gastrointestinal diseases, respectively associated to emesis and diarrhea. In general, both types of foodborne illnesses are relatively mild and usually do last not more than 24 h. Nevertheless, more severe forms have occasionally been reported, including two deaths after the ingestion of food contaminated with high amounts of the emetic toxin and three deaths caused by the diarrheal necrotic enterotoxin CytK1.12,55,88,89 The emetic syndrome has been frequently associated with cereal foods including rice and pasta. Diarrheal toxins have been found in many foods, including milk, vegetables, and meat products.90 The minimal cell density required to provoke both types of diseases is estimated in the range 105–108 CFU/g of ingested food (CFU=colony-forming units). However, there are some reports of emetic syndrome associated with foods containing only 103 CFU/g food.91,92 The diarrheal syndrome is caused by heat-labile hydrophilic enterotoxins produced during the vegetative growth of B. cereus in the small intestine, and is characterized by abdominal pain and diarrhea that are manifested after 8–16 h incubation. Five chromosome-encoded enterotoxins have been characterized: two protein complexes hemolysin BL (HBL) and nonhemolytic enterotoxin (NHE), and three

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enterotoxic proteins: cytotoxin K (CytK), enterotoxin T (BcET) and enterotoxin FM/S (EntMF/S).48,55,93–95 CytK is a 34 kDa hemolytic pore-forming toxin with homology to the β-barrel pore-forming toxins, including staphylococcal β-hemolysin. To date, only three strains containing cytk-1 gene, encoding the more toxic variant of the CytK protein, have been identified: B. cereus strains NVH391/98, NVH883/00, and INRAAF2. Phylogenetic analysis showed that these strains comprise a novel cluster of thermophilic strains genetically distant from the other B. cereus group strains.96,97 Strain NVH 391/98, isolated in 1998 in France at a level of 3×105/g from a vegetable puree, has been implicated in bloody diarrhea that caused the death of three persons.55,98 Recently, the complete genome sequence of this strain revealed the presence of two new important diarrheic toxin operons that may have contributed to the toxicity of this strain in the fatal incident.99 HBL and NHE are tripartite toxins, in which all three components are necessary for maximal cytotoxic activity.100,101 NHE consists of three components with molecular masses of 39, 45, and 105 kDa. HBL contains the protein components B (37.5 kDa), L1 (38.2kDa), and L2 (43.5 kDa).102 A previous study indicated that the cytotoxicity of B. cereus strains was dominated by the amount of secreted NHE and suggested that NHE is the most important toxin that contributes to food poisoning.103 Recently, it has been shown that NHE acts as a pore-forming toxin inducing cell lysis; and that HBL and NHE constitute a superfamily of pore-forming cytotoxins.104 It has been suggested that BcET and EntFM have either an unknown type of enterotoxic action or none at all.105 The emetic syndromes characterized by nausea and vomiting, and induced from 1 to 5 h after consumption of the contaminated food, are associated with the 1.2-kDa cereulide toxin.12,88,93,106 Cereulide concentrations ranging from 0.01 to1.28 µg/g were reported in foods implicated in the emetic type of food poisoning.106,107 The chemical structure and characteristics of this toxin have been studied in detail and recently the molecular basis for its synthesis has been described.91,92,107–109 Cereulide is a hydrophobic, non ribosomally synthesized small cyclic dodecadepsipeptide [(D-OLeu-D-Ala-L-OVal-L-Val)3]. The 24-kb (nt 15094–38668) cereulide synthesis gene cluster (ces) encodes seven proteins involved in the synthesis of the cereulide toxin and is located on a 270 kb plasmid.13,109,110 Cereulide is heat- and proteaseresistant and pH-stable. At low K+ concentration, it acts as an ionophore through mitochondrial membranes and interferes with oxidative phosphorylation.111–114 The diversity of the cereulide producers was investigated by Ehling-Schulz et al.,115 who concluded that, like B. anthracis, the cereulide-producing strains represent a recently emerged virulent emetic clone of B. cereus. These emetic toxin-producing B. cereus strains form a distinct cluster within B. cereus and harbor some specific characters, such as inability to hydrolyze starch and to ferment salicin, poor ability to grow at temperatures below 10°C, growth at 48°C, high heat resistance and low ability of their spores to germinate at 30°C. However, individual cereulide-producing strains differed up to 1000-fold

Molecular Detection of Foodborne Pathogens

in their capacity to produce cereulide.116 B. cereus may be differentiated into 18 serotypes based upon flagellar (H) antigens. Serovar H1 was considered the predominant serovar implicated in the emetic food poisoning syndrome.117 However, a recent study showed that strains of serovars H3 and H12 are also sources of emetic-type food poisoning and that emetic toxin producers belong to two phylogenetically distinct B. cereus clusters.118 Taken together with the discovery of emetic B. weihenstephanensis,4 these data suggest that cereulide-producing strains are progressively diversifying. The fact that the capacity for cereulide production is shared by two phylogenetically distinct B. cereus clusters and by B. weihenstephanensis strains, suggests that the 270 kb plasmid carrying the operon responsible for cereulide production, named pCER270,13,109,110 can undergo to lateral transfer. Indeed, plasmid transfer between members of the B. cereus group has been shown to occur.119–122 The true incidence of B. cereus food poisoning is unknown for a number of reasons. These include the differences in reporting procedures between countries and the relatively short duration of both disease syndromes (<24 h). In addition, the frequency at which single people are affected is usually not monitored. Another reason for this underestimation is the misdiagnosis of the disease, which is symptomatically similar to other types of food poisoning. For example, the symptoms caused by emetic B. cereus resemble those caused by Staphylococcus aureus, while those caused by diarrheal B. cereus resemble those caused by Clostridium perfringens.91,92 In some cases, epidemiological investigations of B. cereus outbreak could not be achieved due to delayed notification, which prevented the acquisition of suspected food samples, or to the lack of a reference laboratory and/or specific diagnostic protocols.123 For instance, between 1991 and 2005, 153 outbreaks were caused by B. cereus in Taiwan, representing 11.2% of the total outbreaks of the period.124 In the USA, from 1982 to 1997, 8781 foodborne-disease outbreaks were reported to the Centers for Disease Control and Prevention (CDC). Of these, 2246 (26%) involved at least five individuals and had sufficient clinical information reported to be considered confirmed foodborne outbreaks. A specific etiology was confirmed by laboratory testing in 697 (31%) outbreaks; among which B. cereus accounted for 1% (10 out of 697 outbreaks with a known etiology).125 Between 1980 and 1997, 2715 cases of B. cereus food poisoning in England and Wales were reported to the Public Health Laboratory Service.126 In Italy and Germany few B. cereus foodborne disease outbreaks have been reported. In June 2000, 173 people presented symptoms of intoxication (nausea and watery diarrhea) after they attended banquettes in Pisa, Italy. A microbiological investigation was performed and HBL-producing B. cereus strains were recovered from stool samples of 19 patients (out of 23 who required hospitalization), foods (more than 102 CFU) and from the rolling board of the confectioner’s shop which was hypothesized to be the source of contamination of all food samples analyzed.127 In 2006, 57 persons (out of 149 participants to a wedding banquet) had a foodborne illness in Salerno, Italy. Ricotta cheese contaminated with B. cereus

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was supposed to be the responsible of this outbreak, although diagnostic certainty was not achieved, as no leftovers were available.123 In June 2006, two emetic food poisoning outbreaks involving 17 children who became sick one hour after eating a rice dish with vegetables, and one student who consumed cooked cauliflower, stored at room temperature for 1.5 days and then reheated, were reported in Germany.128 In France and Belgium, diarrheal and emetic B. cereus fatal food poisoning cases were reported.12,55 In African countries very little is known about the real incidence of B. cereus food poisoning outbreaks, and this could be due to the lack of surveillance data and under-reporting of the disease. In Kenya, 37 food poisoning outbreaks were reported to the Ministry of Health from various parts of the country in the period 1970– 1993. Only 13 of these involving 926 people were confirmed to be due to particular etiological agents, among which B. cereus. However, authors suggested that foodborne disease outbreaks are many more than those recorded by the Ministry of Health due to under-reporting, inadequate investigation of outbreaks, and inadequate diagnostic facilities.129 The type of illness most commonly encountered in a country is influenced by the average diet type and nutrition habits of the population. For example, while in Japan, the emetic illness is reported more frequently than the diarrheal one, in Europe and North America the diarrheal illness is more frequent.112

ingested, in which the presence of B. subtilis could have been a contributing factor. B. subtilis has been associated with emesis following the consumption of large numbers (2×106) of cells.133 Moreover, the detergent surfactin was shown to be toxic toward boar sperm cells through pore formation and destruction of the cell with subsequent loss of mitochondrial activity.134,135 With regard to the other Bacillus species, toxin production was confirmed and in vitro toxicity toward cell lines was demonstrated. For instance, two emetic B. weihenstephanensis strains were able to grow and produce cereulide at 8°C.4 The complete genome sequencing of strain B. weihenstephanensis KBAB4 revealed the presence of a NHE-like toxin on a 400 kb plasmid.99 To our knowledge, up to now there were no reports on food poisoning cases related to these species. However, the risk of food poisoning from psychrotolerant emetic strains in refrigerated foods, and from cyclic lipopeptides-producing strains needs to be further investigated. The loss of cytotoxicity by B. mojavensis following the abolition of surfactin synthesis5 supports a role of this molecule in potential virulence. It has been shown that strains of B. firmus, B. megaterium, B mojavensis, B. fusiformis and B. simplex2,3 can produce heat-stable toxins with physical characteristics and mode of action similar to those of the B. cereus emetic toxin.

10.1.5 Other Foodborne Virulent Bacillus spp.

10.1.6  Detection of B. cereus Toxins

Besides B. cereus, other species in the genus Bacillus have been implicated in food poisoning. Among these, B. weihenstephanensis, B. subtilis, B. licheniformis, B. pumilus, B. mojavensis, B. firmus, B. megaterium, B. simplex, and B. fusiformis have been shown to produce toxins and should be considered of food safety concerns.2,3,5,7,90 In recent years, several authors reported on food poisoning incidents implicating Bacillus species other than the B. cereus: (i) The baby milk powder containing toxigenic B. licheniformis involved in fatal illness of an infant,1 which were later shown to produce the toxin lichenysin A;130 (ii) a B. pumilus-contaminated rice in a Chinese restaurant intoxicated three individuals. Strain NVH891-05 isolated from the contaminated rice grew well at low temperatures (10–15°C) and produced large amounts of a toxic cyclic acylheptapeptide, named pumilacidin.5 Heat-stable toxin-producing B. licheniformis and B. pumilus were also isolated from mastitic milks. The toxins inhibited mobility of boar sperm cells and disturbed the plasma membrane permeability barrier without affecting the mitochondria.131 The B. licheniformis isolates were shown to have ribopattern similar to strains implicated in the infant fatal case of food poisoning, and harbored three lichenysin A synthetase genes lchAA, lchAB, and lchAC;131 (iii) Duca et al.132 reported on a case of infant emesis resulting from consumption of an infant cereal product contaminated with B. cereus and B. subtilis spores. However, the emetic toxin production was not demonstrated and the B. cereus strain was not isolated from the afflicted infants. The emesis has than been attributed to the high bacterial load

The detection of cereulide in suspected contaminated food samples, or in vomit and fecal samples from ill patients or in the isolates obtained from those samples, represents a necessary clue to confirm B. cereus as the causative agent. At present, different tests exist and are commonly used for the detection of diarrheal toxins like the vascular permeability reaction (VPR), the ligated rabbit ileal loop, and cell cytotoxicity assays. VPR consists in injecting cell-free culture supernatants (0.1 ml) intradermally into rabbits weighting 2.5–3.0 kg. After 3 h, 4 ml of 2% Evans blue dye solution is injected intravenously. Measurements of perpendicular diameters of the zones of light and dark blue, and necrosis when present, are made after 1 h.136 The ligated rabbit ileal loop assay consists in the injection of the material to be tested (e.g., the bacterial culture supernatant) into 5 cm ileal loop of female New Zealand white rabbits. The reaction is considered positive if the ratio of the volume of fluid accumulation to loop length is >0.5.94 In the cytotoxicity assays filtered supernatant is supplemented to a cell line and the effects of the treatment on the cells is evaluated. A number of cell lines are susceptible to the diarrheal toxins. The most commonly used are Vero (monkey kidney) and Chinese hamster ovary (CHO) cell lines, although other cell lines have been used including HeLa S3, Human Embryonic Lung (HEL) and McCoy cell lines.136,137 Gray et al.90 developed an innovative assay using a murine hybridoma Ped-2E9 cell model. Culture supernatants containing enterotoxins are added to a Ped-2E9 cell line and analyzed for cytotoxicity with an alkaline phosphatase release assay. The assay was shown to be rapid (results

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obtained within 1 h) and to have 25- to 58-fold-higher sensitivity than the CHO assay. Although cell cytotoxicity assays are an inexpensive and convenient method for diarrheal toxin detection, they present many drawbacks. B. cereus produces many extracellular virulence factors such as phospholipases and sphingomyelinase, which may be cytotoxic to the cell lines.38,138,139 The test is time-consuming since it needs an enrichment step and requires the constant maintenance of cell lines. Commercial kits have been developed for the detection of the diarrheal toxins of B. cereus. Two immunological assays are commercially available: the B. cereus Enterotoxin Reverse Passive Latex Agglutination (BCET-RPLA) kit (Oxoid, Basingstoke, UK) and the Tecra Bacillus Diarrheal Enterotoxin Visual Immunoassay (BDE-VIA) kit (Tecra Diagnostics, Frenchs Forest, Australia) for detection of the L2 component of HBL (HblC) and the 41-kDa subunit of NHE, respectively. Both kits require the culture of B. cereus isolates for 6–18 h prior to testing. The culture supernatants are then tested for the enterotoxin-related proteins. To our knowledge, commercial kits are not yet available for the detection emetic toxin of B. cereus. Cereulide is traditionally assessed using a HEp-2 tissue culture assay by observing vacuole formation106,140,141 or colorimetrically, by using the 3-(4,5-dimethylthiazolyl-2)-2,5-diphenyltetrazolium bromide (MTT) metabolic staining assay.142,143 An assay based on the uncoupling of respiratory activity of rat liver mitochondria has been developed for emetic toxin.112 Another bioassay, based on the loss of motility of boar spermatozoa, has also been developed.144,145 However, this assay has been found to be not enough specific, since other toxins were shown to affect sperm motility.146 Recently, a Computer Aided Semen Analysis (CASA) study of the boar semen motility has been demonstrated to be an appropriate assay for detection of cereulide, which induces motility ceasing at concentrations lower than 20 ng/ml, in comparison with other toxins that have the same effect at much higher concentrations. Since the assay detected cereulide indirectly, the presence of cereulide in positive samples was verified by HPLC-MS.147–149

10.1.7 Detection of B. cereus Cells in Food and Environmental Samples Standard isolation and enumeration of B. cereus from foods, the environment and clinical settings are generally performed by using conventional selective plating media. Two standard media are recommended by food authorities: polymyxinegg yolk-mannitol-bromothymol blue (PEMBA, Oxoid) or mannitol-egg yolk-polymyxin (MEYP, Oxoid) agar.150 When grown on PEMBA, B. cereus produces typical crenated colonies that retain the turquoise blue of the pH indicator (bromothymol blue) due to their inability to produce acid via mannitol fermentation. A zone of egg-yolk precipitation is produced through lecithinase activity, which involves the cleaving of lecithin (phosphatidylcholine) into phosphorylcholine and diglyceride. Polymyxin B is used in the media as a selective agent to suppress Gram-negative bacteria.151

Molecular Detection of Foodborne Pathogens

Recently, two new chromogenic plating media, B. cereus group plating medium (BCM, Biosynth AG, Switzerland) and Chromogenic B. cereus agar (CBC; Oxoid) were developed based on the activity of two different hydrolyzing enzymes: the phosphatidyl inositol phospholipase C (PI-PLC) for BCM, and the β-D-glucosidase for CBC. On BCM, 5-bromo-4chloro-3-indoxylmyoinositol-1-phosphate is cleaved by PIPLC and gives to B. cereus colonies a homogeneous blueturquoise color. The colonies are sometimes surrounded by a blue halo. CBC contains 5-bromo-4-chloro-3-indolylβ-glucopyranoside that is cleaved by β-D-glucosidase and results in white colonies with a blue-green centre. However, in addition to being time-consuming and unable to indicate the toxin production capability of the isolates, the performance of these culture-dependent detection methods are debated. In fact, some B. cereus strains, especially highly toxic strains with atypical growth characteristics, lack one or more of the key characteristics on these media, a feature that could lead to their misidentification.152 In addition, presumptive B. cereus isolates must then be tested by several biochemical and microscopic procedures to confirm whether they are true B. cereus, due to the phylogenetically close relationship between the different species. Crystal formation is one key test that positively identifies B. thuringiensis, while rhizoid colony morphology is a phenotypic character of the species B. mycoides and B. pseudomycoides.27 Thus, acrystalliferous variants of B. thuringiensis or nonrhizoid variants of B. mycoides/B. pseudomycoides may be misidentified as B. cereus. Moreover, the difficulty in clearly identifying isolates within the B. cereus group is also increased by the presence of enterotoxin-producing B. thuringiensis strains (Raddadi et al., unpublished data).153–156 Blood-agar medium could also be used to select HBL-producing B. cereus strains that show discontinuous hemolytic patterns.157 Based on monoclonal antibodies that detect specifically the B component of HBL and the nheA component of NHE, Moravek et al.158 described a colony immunoblot assay (CIA) that enables the identification of HBL and NHE-producing B. cereus isolates grown on blood agar within 24 h. Over years, several biosensor-based techniques for the detection of foodborne pathogens and bioterrorism agents have been developed and reported in the literature. A biosensor is an analytical device that integrates a biological sensing element with an electrical transducer to quantify a biological event into an electrical output. Recently, Pal et al.159 used immunochemical techniques for the development of a B. cereus biosensor. The approach utilizes the concept of a direct charge transfer (DCT) of electrons in a voltage controlled switch format for bacterial detection. The biosensor uses rabbit polyclonal anti-B. cereus antibodies as the biological sensing element and polyaniline nanowires as the electrical transducer. The biosensor was shown to have high sensitivity, being able to detect the presence of B. cereus at concentrations as low as 101 CFU/ml, and rapid with a detection time being only 6 min. In addition to these different detection methods, molecular detection systems for foodborne pathogenic bacteria in

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general, as well as for diarrheal and emetic B. cereus strains, have been developed along the years. These include PCRs (conventional, multiplex, and real-time) and oligonucleotide DNA microarrays. Jin et al.160 developed DNA microarrays for the detection and identification of intestinal pathogens using two universal PCR primers to amplify two variable regions of bacterial 16S and 23S ribosomal RNA genes. Two oligonucleotide microarrays based on 16S rRNA gene sequences of predominant human intestinal bacterial species were reported for the detection of intestinal bacteria in fecal samples collected from human subjects, and for the detection of foodborne pathogenic bacteria, respectively.161,162 However, these tools could not be useful for the discrimination among the bacteria of the B. cereus group due to the close phylogenetic relationship between the different species that harbor mostly 16S rRNA gene sequences. Recently, Park et al.163 developed a multiplex PCR that targets gyrB and groEL genes as diagnostic markers for the simultaneous detection and identification of the B. cereus group bacteria from food samples. A 400 bp PCR fragment was amplified from the groEL gene for all the bacteria of the B. cereus group, while amplicons of 253, 475, 299, and 604 bp were yielded for the gyrB gene from B. anthracis, B. cereus, B. thuringiensis, and B. mycoides, respectively. PCR has been extensively used to amplify all the diarrheal toxin-producing genes,13,89,127,154,164 and genes encoding the emetic toxin cereulide, after their identification and sequencing.91,92,108,115 However, the presence of a toxin gene does not necessarily indicate that the bacterium is capable of producing the protein in concentrations high enough to determine the disease. For example, different emetic B. cereus strains were shown to have 1000-fold difference in their capacity to express cereulide.116 Another major drawback of conventional PCR is the requirement for post-PCR analysis by gel or capillary electrophoresis that is time-consuming, and bears the risk of false-positive results due to laboratory contamination. Thus, introduction of more rapid and sensitive detection systems, such as quantitative real-time PCR (RT-PCR), is required. In principle, two different chemistries are available for real-time detection of PCR products: fluorescent probes that bind specifically to certain DNA sequences (Taq Man-based RT-PCR) and fluorescent dyes that intercalate in any double-stranded DNA (SYBR green I-based RT-PCR). Based on the sequence of the cereulide synthetase (ces) gene, Fricker et al.128 developed novel diagnostic assays that were applied successfully to identify the causative agent of two recent emetic food-poisoning outbreaks in Germany. The methods developed were a Taq Man-based RT-PCR assay targeting a highly specific sequence of the ces gene, and a duplex SYBR green I-based RT-PCR assay for onestep differentiation between emetic B. cereus and S. aureus. Yang et al.124 developed a SYBR green-based RT-PCR assay for the rapid (analysis is performed in less than 3 h) quantitative detection of B. cereus in food (cooked rice, milk, and chicken meat) and fecal samples, using the nheB gene as target. Combined with the most probable number (MPN) technique (MPN-RT-PCR assay) for the enumeration of low

loads of contamination, the assay was performed in less than 8 h and the detection limit was as low as 1 CFU/ml. The MPN-RT-PCR assay could be used as an alternative method for detecting low levels of contamination. It provides stronger quantification than the traditional enrichment step, allows the dilution of PCR inhibitors and the discrimination of viable and dead cells. In fact, when applying RT-PCR to the detection of pathogens in foods without enrichment, the method usually consistently detects 102–103 CFU or more per gram of sample.124,128,165 This decrease in the sensitivity of RT-PCR, which theoretically can reach one copy of target gene detected per reaction, is mainly due to PCR inhibitors such as lipids, proteases, and divalent cations present in dairy products.166,167 To overcome this problem and enhance the sensitivity of the detection, most available detection systems require selective enrichment steps, especially for low pathogen concentrations. However, this could prevent the appreciation of original pathogen contamination levels. Recently, Fukushima et al.165 developed a method for the rapid separation and concentration of B. cereus cells (and also other foodborne pathogenic bacteria) from food matrices that can be used prior to real-time quantitative PCR and viable-cell counting. Using these combined methods, the target organisms in the food samples can be concentrated up to 250-fold and detected at cell concentrations as low as 101 CFU/g. In the following paragraphs, we will report on procedures that can be adopted for the detection and identification of B. cereus using modern molecular methods. Principles and procedures, from sample preparation to application of detection techniques, are illustrated and discussed.

10.2  Methods 10.2.1 Sample Preparation Bacterial isolation and enumeration from food samples remain an integral component in the sample preparation prior to molecular confirmation of foodborne pathogens concerned. Additionally, bacterial DNA can be extracted directly from food matrices both for microbiological quality evaluation and risk assessment studies of foodborne pathogens of interest, and for epidemiological diagnosis of food poisoning illnesses using molecular detection techniques. In the latter case, the incriminated bacterium is also isolated on selective medium and further identified by biochemical and/ or molecular methods. 10.2.1.1  B. cereus Isolation and Enumeration To enumerate B. cereus spores, 25 g or 25 ml of food samples are homogenized with 225 ml of peptone saline solution (PSS) containing 8.5 g/l NaCl and 1 g/l of neutralized bacteriological peptone. Ten milliliters of the homogenate (diluted 1:10) are transferred into sterile tubes, incubated at 80°C for 10 min, and then cooled in melting ice prior to further serial dilutions in PSS, until 1:1000 dilution. In order to detect and enumerate low levels of B. cereus spores, a three-tube MPN procedure could be performed using as growth medium

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tryptone soya broth (TSB, Oxoid) supplemented with 100 mg/l of polymyxin B sulfate. After incubation at 30°C for 24 h, tubes are examined visually for turbidity. A loopful of culture from positive MPN tubes is streaked onto one of the chromogenic B. cereus group selective media (MEYP; BCM; CBC) plates and incubated at 30°C for 24–48 h. Colonies with a typical B. cereus morphotype are purified on tryptone soya agar (TSA; Oxoid). In order to detect and enumerate high levels of spores of B. cereus, the spread-plate procedure is carried out as follows: 1 ml volume of the 1:1000 dilution is spread plated in duplicate onto one of the three B. cereus group selective media agar plates and incubated at 30°C for 24–48 h. The plates with 15–150 typical B. cereus colonies are counted. One to five randomly selected typical colonies are purified. The total counts of B. cereus spores are based on percentage of colonies tested that were confirmed as B. cereus, and expressed as CFU/g of sample. The purified isolates are stored in nutrient broth (Oxoid) with 20% glycerol at –80°C until use.84,162 10.2.1.2 DNA Isolation DNA template for PCR reactions can be prepared from the purified isolates by a simple boiling method as following. Cells are collected by centrifugation (5000×g for 2 min) from 1 ml of a 4-h culture grown in nutrient broth at 30°C. The cell pellet is washed once with 500 µl of sterile MilliQ water or TE buffer (pH=8), centrifuged, resuspended in100 µl of sterile MilliQ water or TE buffer and boiled for 10–15 min. The boiled samples are centrifuged at 10000×g for 5 min to precipitate cell debris and the supernatants, which contain nucleic acids, are collected and stored at –20°C until use. There are also several commercial kits available for total DNA extraction from overnight grown pure bacterial cultures such as AquaPure genomic DNA isolation kit (BioRad) or QIAamp DNA stool mini kit (QIAGEN). The phenolchloroform extraction protocol is also useful to obtain pure high molecular weight genomic DNA.168 Total DNA from food samples can be isolated using the AquaPure genomic DNA isolation kit (Bio-Rad), the QIAamp DNA Mini Kit (Qiagen), the Wizard DNA Purification resinbased kit (Promega), the NucleoSpin food kit (MachereyNagel), or a simple boiling method. Yang et al.124 extracted DNA using the QIAamp DNA stool mini kit (Qiagen) with some modifications to the manufacturer’s instructions. For pure cultures, bacteria are placed in brain heart infusion broth and incubated overnight at 30°C. DNA is extracted from 200 µl of each individual culture. For cultures in spiked food or fecal samples, 200 µl or 200 µg of spiked samples are added to 1.4 ml of buffer ASL and heated for 15 min at 99°C. After heating, the manufacturer’s steps are followed up to the proteinase K step. At this time, 30 µl of proteinase K (20 mg/ ml) is placed into a new microcentrifuge tube, and 400 µl of the supernatant obtained from “Inhibit Extreatment” step is added, followed by 400 µl of buffer Al and 400 µl of ethanol. Since the total volume of the resulting lysate exceeds the volume of the spin column, the lysate is applied twice to the

Molecular Detection of Foodborne Pathogens

spin column. Finally, after the washing steps 40 µl of buffer AE is added to elute the DNA.

10.2.2  Detection Procedures The traditional methods for strain identification, looking to metabolism and physiology, can be time consuming and laborious. As well, the culture-dependent techniques for direct isolation and identification of B. cereus suffer the same problems. In addition, these techniques do not allow the differentiation among the B. cereus group isolates due to the close phylogenetic relationship between the species in this group and the presence of borderline isolates between B. cereus and B. anthracis. This could be of high risk from a human safety point of view. In fact, although B. cereus has been implicated in some fatal cases, the illnesses caused by this pathogen are usually mild. On the contrary, up to 40% of people having gastrointestinal anthrax die after antibiotic treatment.21 Although rare (less than 1% of reported anthrax cases), foodborne anthrax cases have been reported after ingestion of B. anthracis-contaminated meat.21 The gastrointestinal anthrax disease is characterized by fever, nausea, vomiting, abdominal pain, and bloody diarrhea,19,22 symptoms that are somehow similar to intoxication with B. cereus. On the other hand, while B. anthracis could be differentiated from B. cereus based on biochemical traits, a problem still exists for the B. cereus/B. thuringiensis borderline strains, which can display disease symptoms similar to B. anthracis. Recently a strain isolated from humans, identified as B. cereus on the basis of phenotypic and molecular data, caused symptoms similar to those of the anthrax disease and it was shown to harbor a virulence plasmid very similar to pXO1, while a capsule plasmid completely different from pXO2.60 These isolates could be discriminated from B. anthracis by different molecular techniques such as RT-PCR assays that target the nonsense mutation in the pleiotrophic regulator gene plcR.53,169–171 However, the approach requires expensive RT-PCR platforms, limiting equipment in most laboratories in developing countries where anthrax is endemic.67 10.2.2.1 Detection of B. anthracis by RSI-PCR Principle: Gierczyn´ ski et al.172 developed a simple and cost effective RSI-PCR-based assay as an alternative to this RT-PCR approach.36 The assay precisely detects the B. anthracis-specific nonsense mutation in plcR gene by restriction digestion with the endonuclease SspI. The requirements of this assay are limited to standard PCR equipment and an agarose or a polyacrylamide gel electrophoresis (PAGE) apparatus, that makes it available for most routine diagnostic laboratories worldwide. The nonsense mutation in the B. anthracis gene plcR generates a termination codon (UAA) in place of glutamic acid (GAA) in position 209 of the amino acid sequence of the functional PlcR.53 In order to detect thymidine (T), the key nucleotide in the plcR nonsense mutation of B. anthracis, an artificial SspI specific cleavage site can be introduced into a PCR

Bacillus

product generated using the 60-bp long AplR primer (5′-AT GTCATACTATTAATTTGACACGATAGTTCAATAGCT TTATTTGCATGACAAAGCGAAT-3′), modified at the 3′ end to specifically introduce the SspI cleavage site in the amplified plcR fragment. At positions 58 and 59 of the AplR primer, two adenine (A) bases determine, when the amplicon is generated, the incorporation in the PCR product of an artificial cleavage site (AATATT) for SspI only in the plcR amplicon deriving from B. anthracis. These two adenines replaced cytosine (C) and thymine (T), which complement G and A at positions 1147 and 1146 (positions referred to sequence accession number AF132086), respectively, of the truncated plcR of B. anthracis strain 9131.53 Primer AplF (5′-GCTCAATCAACAATTGGCAGG-3′) can be used in combination with primer AplR for the amplification of a 278 bp fragment of the plcR gene.172 Procedure: (1) Prepare PCR mixture (25 µl) containing 1× PCR buffer (10 mM Tris-HCl pH 8.9, 2 mM MgCl2, 50 mM KCl), 10 pmol of each primer (AplF and AplR), 0.2 mM of each dNTPs and 5–10 ng of genomic template DNA. (2) Carry out a hot start step at 95°C for 5 min in a PCR cycler before adding Taq DNA polymerase (to avoid formation of primer dimers whose formation could be favored by the 60-bp forward primer used). (3) Add 1 U of Taq DNA polymerase (Polgen), and perform the cycling program consisting of an initial denaturation step at 94°C for 3 min, 35 cycles of denaturation at 94°C for 40 sec, annealing at 58°C for 30 sec and extension at 72°C for 30 sec and a final extension at 72°C for 5 min. (4) Transfer 6 µl of PCR product in a tube containing 5–7 U of restriction enzyme SspI with appropriate reaction buffer (total volume 20 µl) and incubate at specified temperature and time. (5) Separate the digestion products in 4% high resolution agarose (MP Biomedicals) in TAE (40 mM Tris-acetate pH 8.0, 1 mM EDTA) at a constant voltage of 6 V/cm for 3 h. (6) Visualize the specific DNA fragments (about 60 and 220 bp for B. anthracis; 120 and 158 bp for some B. thuringiensis strains that have native SspI restriction site in the plcR gene) with conventional ethidium bromide staining. Note: it is also possible to separate the digestion products onto 8% nondenaturing polyacrylamide gel (Applichem) using a conventional chamber for vertical electrophoresis. In this case, DNA is visualized by either ethidium bromide staining or silver staining. Further details of the procedures to be adopted for RSI-PCR analysis can be found in Daffonchio et al.,36 Daffonchio et al.,43 and Gierczyn´ ski et al.172 This test was applied to 47 strains of B. anthracis that were isolated from four different countries (France, Georgia, Poland, and Russia), including

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vaccinal strains, and all gave a SspI restriction profile with two fragments of about 60 and 220 bp as expected.172 It is important to notice that some B. thuringiensis strains gave aspecific SspI restriction profile of about 120 and 160 bp. However, these strains were shown to have a native SspI restriction site that splits PCR product into fragments of 120 and 158 bp. In addition, some B. cereus tested in this assay gave a plcR amplified fragment of the expected length (290 bp) but was not cleaved by the restriction enzyme.172 10.2.2.2 Detection of B. cereus Toxin Genes by Multiplex PCR With regard to B. cereus toxins, two commercial kits are available for the detection of the diarrheal HBL and NHE (see Section 10.1.4), while no kits are actually available for the detection of the emetic toxin cereulide, due to its low antigenic characteristics,173 or the necrotic toxin CytK1. Over the years, to allow the rapid detection of the genetic determinants of these toxins, different molecular methods have been developed, including microarray 174 conventional PCR and RT-PCR. Principle: Ngamwongsatit et al.156 developed a multiplex PCR method using new specific PCR primers for detection of eight enterotoxin genes (hblCDA, nheABC, cytK, and entFM) in a single PCR reaction. Procedure: (1) Prepare PCR mixture (in a final volume of 20 µl) containing 5 µl of template DNA prepared by the boiling method, 1× PCR buffer (10 mM Tris-HCl pH 8.3 and 50 mM KCl), 1.5 mM MgCl2, 200 mM of each dNTP, 0.2–0.4 mM of primers (targeting hblD, nheB, hblA, nheA, hblC, nheC, cytK, and entFM genes) and 5 U of Taq DNA polymerase. (2) Carry out the PCR amplification in cycler with an initial denaturation at 95°C for 5 min, 30 cycles of 94°C for 45 sec, 54°C for 1 min, 72°C for 2 min and a final extension at 72°C for 5 min. (3) Separate the amplicons on 1.5% agarose gels and visualize the fragments of predicted size of 1018, 935, 884, 759, 695, 618, 565, and 486 bp for hblD, nheB, hblA, nheA, hblC, nheC, cytK, and entFM genes, respectively, from B. cereus/B. thuringiensis. Note: by using this assay, Ngamwongsatit et al.156 categorized the 616 isolates into four groups. All eight genes were detected in group I. Group II and III lacked hblCDA and cytK, respectively, while in group IV, both hblCDA and cytK were missing. However, the applicability of the method was not tested on food samples, which often contain PCR inhibitors. This multiplex PCR is an informative and easy-to-handle tool for the detection of enterotoxin genes in individual isolates purified from food samples. However its efficiency for the direct

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detection of toxin-producing B. cereus/B. thuringiensis in food samples should be tested to be used as a tool for risk assessment. 10.2.2.3 Detection of B. cereus Toxin Genes by RT-PCR Principle: Yang et al.124 developed a SYBR green-based RT-PCR for the detection of B. cereus group cells in different food and fecal samples. They choose the nhe gene that codify for the diarrheal toxin NHE, because this gene was shown to be present in 100% of B. cereus and B. thuringiensis strains.156 Another reason of the choice is related to the fact that NHE toxin is the most important toxin in B. cereus diarrheal food poisoning.103 Procedure: (1) Prepare RT-PCR mixture (25 µl) containing 12.5 µl of SYBR Premix Ex Taq (2×) (TaKaRa Bio Inc.), 0.5 µl of ROX reference dye II (50×), 10 µM of each primer and 2.5 µl of template DNA extracted by boiling, and two primer pairs (consensus primers: SG-F3, 5′-GCACTTATGGCAGTATTTGCAGC-3′, and SG-R3,  5′-GCATCTTTTAAGCCTTCTGGTC-3′; degenerate primers: mSG-F3, 5′-GCACTKATGGC AGTATTTRCR GC-3′, and mSG-R3, 5′-GCATCTT TYARGCCTTCTGGTC-3′) in order to overcome false-negative results due to the sequence polymorphisms characterizing this gene. (2) Perform PCR amplification on a cycler (e.g., an Mx3000P apparatus, Stratagene), with the thermal protocol consisting of in initial denaturation at 95°C for 30 sec, followed by 40 cycles at 95°C for 5 sec and 60°C for 30 sec. (3) Examine the amplified products by gel electrophoresis to confirm that the results reported by the Mx3000P correlated with the amplification of a product with the expected size (152 bp). Note: the performance of the two primer pairs was evaluated by Yang et al.124 with 10 ng of purified DNA from 60 strains of the B. cereus group and was shown to be specific for bacteria of this group. This method was shown to be highly sensitive detecting as less as 102–107 CFU/g of B. cereus in cooked rice and chicken samples, and thus was supposed to be sufficient to be used as a quick and routine technique for the quantification of B. cereus group cells. However, a problem still exists for the detection of low-level contamination. In this case, an enrichment step could be incorporated in order to allow bacterial pathogens to multiply and reach detectable levels. However, this kind of method prevents the quantification of original contamination levels. In order to solve the problem of low-level contamination detection, the RT-PCR can be coupled to a three-tube MPN assay. This method was applied to 30 food samples, and was shown to have a detection

limit of as low as 1 B. cereus CFU/ml, which is more sensitive than an RT-PCR assay without enrichment.124 This MPN RT-PCR assay provides stronger quantification than the traditional enrichment and is important for the discrimination of viable and dead cells. This method is useful for detection and quantification of cells of the B. cereus group and not only of the B. cereus species, since nhe gene is widely distributed among bacteria in this group. 10.2.2.4 Detection of Emetic B. cereus Strains by RT-PCR The first conventional PCR-based assay for the detection of emetic B. cereus strains has been described in 2004.91,92 Primers EM1F (5′-GACAAGAGAAATTTCTACGAGCAA GTACAAT-3′) and EM1R (5′-GCAGCCTTCCAATTACTC CTTCTGCCACAGT-3′) amplify a fragment of 635 bp from emetic B. cereus genomic DNA. The specificity of the assay was assessed using a panel of 178 bacterial strains; neither false-positive nor false-negative signals were detected.91,92 Principle: recently, after the biosynthetic pathway for nonribosomal synthesis of cereulide and the related genes have been deciphered,115,176 a TaqMan-based as well as a SYBR green I-based duplex RT-PCR assays that target a highly specific part of the cereulide synthetase (ces: accession no. DQ360825) genes were developed by Fricker et al.128 for the detection of emetic B. cereus and its differentiation from S. aureus. The target probe for emetic B. cereus was labeled at the 5′ end with the reporter dye 6-carboxyfluorescein (FAM), and the IAC probe was labeled at the 5′ end with 5-hexachloro-6-carboxyfluorescein (HEX). Both probes were labeled at the 3′ ends with tetramethyl-6-carboxyrhodamine (TAMRA). Procedures: (i) TaqMan-based RT-PCR: (1) Prepare PCR mixture (25-µl) containing 12.5 µl Brilliant Q PCR Multiplex Mastermix (Stratagene), 0.5 µM of each primer (ces_TaqMan_ for: 5′-CGCCGAAAGTGATTATACCAA-3′; ces_TaqMan_rev: 5′-TATGCCCCGTTCTCAA ACTG-3′; IAC_for: 5′-GCAGCCACTGGTAA CAGGAT-3′; IAC_rev: 5′-GCAGAGCGCAGAT ACCAAAT-3′), 0.2 µM of each probe (ces_ TaqMa n _ probe:  5′-FA M- G G GA A A ATA ACGAGAAATGCA-TAMRA-3′; IAC_probe: 5′-HEX-AGAGCGAGGTATGTAGGCGGTAMRA-3′), approximately 170 copies of plasmid DNA pUC19 (Fermentas), and 5 µl of DNA template. Controls containing 5 µl of only TrisEDTA buffer were included in each run to detect any contamination. (2) Carry out the amplification in a Stratagene MX3000P PCR system (Stratagene) with the thermal cycling consisting of 95°C for 10 min followed by 40 cycles at 95°C for 15 sec and

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Bacillus

55°C for 60 sec. The maximum ramp rate of the Stratagene MX3000P RT-PCR system was 2.5°C/sec. (ii) SYBR green I RT-PCR: (1) Prepare PCR mixture (25-µl) containing 12.5 µl SYBR Premix Ex Taq (Takara Bio, Inc.), 1–5 µl of template DNA (depending on extraction method), 0.3 µM for each B. cereus primer (ces_SYBR_F: 5′-CACGCCGAAAGTGATTATACCAA-3′; ces_SYBR_R: 5′-CACGATAAAACCACTGA GATAGTG-3′), and 0.12 µM for each S. aureus primer (sa_SYBR_F: 5′-CGTGTTGA ACGTGGTCAAATCA-3′; sa_SYBR_R: 5′-CA CCTTCGTCTTTTGATAATACG-3′). (2) Perform PCR amplification in a Stratagene MX3000P RT-PCR system with the following cycling program: one cycle of 95°C for 10 sec, and 45 cycles at 95°C for 10 sec and 60°C for 30 sec. Note: the same conditions can be used with the Smart Cycler II system (Cepheid), but with the ramp rate set on maximum (10°C/sec) for the simplex detection of emetic B. cereus strains and altered to 3°C/sec for simplex and duplex detection involving the primers specific for S. aureus. The real-time assays developed were shown to detect 101–103 CFU/g of emetic B. cereus directly from food samples without the need for further enrichment steps. Lower cell numbers (one B. cereus CFU/g) can be detected after a short enrichment time (4–6 h). After 6 h of enrichment, it was possible to detect one B. cereus CFU/g and 103 CFU S. aureus/g of food by the SYBR green duplex assay when emetic B. cereus was present in the same enrichment. Foods incriminated in emetic outbreaks were reported to have 105–108 CFU B.cereus/g or between 105 and 106 S. aureus CFU/g.177 Thus samples from emetic food poisonings can be processed and analyzed within short time (1.5–6 h). However, once the presence of B. cereus is detected by this assay, a further analysis of the emetic toxin cereulide is still needed due to its heat stability, especially in the cases of reheated foods. Three methods for detection of the emetic toxin have been described including a cytotoxicity assay using HEp2 or CHO cells, HPLC-MS analysis, and a spermbased bioassay. The cytotoxicity assay could be performed from pure B. cereus isolates as following: cells are grown in 20 ml of skim milk medium for 18 h, and, after autoclaving, an aliquot of the preparation is serially diluted (two-fold) in 96-well plates by using Earle’s minimal essential cell culture medium supplemented with 1% fetal calf serum, 1% (vol/vol) sodium pyruvate (100 mM), 2% (vol/vol) L-glutamine (200 mM), 0.2% (vol/vol) penicillin-streptomycin (10,000 U/ml), and 2% ethanol as a diluent. Immediately after this, HEp-2 cells (0.15 ml; 105 cells per well) are added, and the plates are incubated for 48 h at 37°C in a 5% CO2 atmosphere. Toxicity titers are determined by using the cell proliferation

reagent WST-1.103,175 The cytotoxicity assay could also be performed directly from food samples. In this case, 5 g (or 5 ml) of food sample are homogenized in 5 ml sterile Milli-Q water and autoclaved (20 min at 121°C). Ten microliters of the cell-free supernatant are then tested in a HEp-2 cell culture assay as described above. Both cytotoxicity assay and sperm-based bioassay are semi-quantitative, and the conclusive identification of cereulide is only provided by HPLC-MS analysis.146,147

10.3 Conclusions and Future Perspectives It is virtually impossible to ensure that foods are free from spore forming bacteria such as the ubiquitous bacterium B. cereus. In most food poisoning cases, illness is associated with the heating of precooked food held for too long at inappropriate storage temperature. However, if food is cooked and stored correctly, B. cereus should not constitute serious problems. Toxin-producing Bacillus spp. seem to be rare among isolates from water, food, and food environments, and none of the toxins detected are similar to the B. cereus enterotoxins or to cereulide. However, although rare, Bacillus spp. outside the B. cereus group might still be involved in food poisoning through foods that are considered safe by the public. Thus, it is interesting to develop detection methods of Bacillus spp. in food ingredients and in the food production plants. Several molecular detection methods are actually available for B. cereus and its toxins. These include conventional PCR, multiplex PCR, and RT-PCR. However, although highly specific and sensitive, these molecular methods have some drawbacks. For example, the conventional PCR could be used for qualitative detection of the presence of the foodborne pathogenic bacterium, but it is not usable for risk assessment evaluation since further confirmation methods, such as DNA hybridization or sequencing, are always needed. This method bears also the risk of false-positive results due to laboratory contamination. The multiplex PCR could be a good alternative to detect different pathogens or different toxins of the same pathogenic bacterial species simultaneously. However, in same cases it is possible to get false negative results due to the presence of different primer pairs in the same reaction mix, or due to different annealing temperatures. In addition, this method is qualitative and do not allow quantification of the pathogenic bacterium or of its toxins. In order to quantify the presence of a specific pathogenic bacterium or its toxins, TaqMan or SYBR green-based RT-PCRs could be applied. However, although highly sensitive, this technique could be biased by the presence of PCR inhibitors in complex matrices (such as foods and fecal samples) and cannot discriminate between DNA originating from alive and dead cells. This method allows the detection of 102–104 CFU/g of sample and thus lower levels of the contaminants cannot be detected, requiring an enrichment step. Enrichment steps are time-consuming and could overestimate the initial level of contamination. Another drawback of RT-PCR is the necessity of expensive and sophisticated real-time platforms and

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reagents that cannot be available in most laboratories especially in developing countries. The confirmation of B. cereus as the causative agent responsible for foodborne disease is dependent upon a combination of different clues like food consumption history, nature of the symptoms and detection of the bacterium in the implicated food and/or the patient vomit or feces. It is also necessary to demonstrate either that the same serotype isolated from the vomit/feces is also present in the implicated food, or that the isolate is toxigenic. The rapid detection of microbes in food samples is becoming more critical and the development of rapid and sensitive methods is of great interest for human safety. Molecular techniques can be used to confirm the identity and the nature of B. cereus isolates. These include multiplex PCR that allows the detection of the different enterotoxin genes and RT-PCR for both diarrheal and emetic toxin genes. However, there are still some limitations in the application of these methods. For example, the DNA template used for PCR amplification of the genes does not necessary derive from viable cell and thus also dead bacterial cells are quantified by the RT-PCR. Also, the amplification of the genes by PCR could not necessary be a confirmation of the incrimination of a B. cereus strain as a food poisoning agent because the capability of the B. cereus isolates to express emetic toxin genes was shown to vary 1000-fold between the different strains. In addition, several studies showed that, although the diarrheal toxin genes were amplified by PCR from different strains, their expression, or at least the expression of all the subunits of the three component toxins was not always confirmed. On the other hand, it is important to take into account the presence of PCR inhibitors in food matrices which could lead to false-negative results, and that the sensitivity of quantitative RT-PCR is affected by the presence of DNA polymorphisms of the toxin genes. In conclusion, from a food safety perspective, important foodborne pathogens within the genus Bacillus are the species of the B. cereus group and in particular B. cereus. However, other species within the genus should be considered when examining suspected samples, since several strains of Bacillus species other than those of the B. cereus group have been described to produce lipophilic compounds such as surfactants with potential toxicity for humans. As far as the detection of B. cereus, a combination of molecular methods, including PCR amplification of the toxin-encoding genes and cytotoxicity assays represent the major tools necessary for risk assessment evaluation.

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Molecular Detection of Foodborne Pathogens 163. Park, S.H. et al. Simultaneous detection and identification of Bacillus cereus group bacteria using multiplex PCR. J. Microbiol. Biotechnol., 17, 1177, 2007. 164. Jackson, S.G. Rapid screening test for enterotoxin-producing Bacillus cereus. J. Clin. Microbiol., 31, 972, 1993. 165. Fukushima, H. et al. Rapid separation and concentration of food-borne pathogens in food samples prior to quantification by viable-cell counting and real-time PCR. Appl. Environ. Microbiol., 73, 92, 2007. 166. Feng, P. Impact of molecular biology on the detection of foodborne pathogens. Mol. Biotechnol., 7, 267, 1997. 167. McKillip, J.L. and Drake, M. Real-time nucleic acid–based detection methods for pathogenic bacteria in food. J. Food Prot., 67, 823, 2004. 168. Sambrook, J., Fritsch, E.F. and Maniatis, T. Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. 1989. 169. Slamti, L. et al. Distinct mutations in PlcR explain why some strains of the Bacillus cereus group are nonhemolytic. J. Bacteriol., 186, 3531, 2004. 170. Easterday, W.R. et al. Specific detection of Bacillus anthracis using a TaqMan mismatch amplification mutation assay. Biotechniques, 38, 731, 2005. 171. Easterday, W.R. et al. Use of single nucleotide polymorphisms in the plcR gene for specific identification of Bacillus anthracis. J. Clin. Microbiol., 43, 1995, 2005. 172. Gierczyn´ ski, R. et al. Specific Bacillus anthracis identification by a plcR–targeted restriction site insertion-PCR (RSIPCR) assay. FEMS Microbiol. Lett., 272, 55, 2007. 173. Melling, J. and Capel, B.J. Characteristics of Bacillus cereus emetic toxin. FEMS Microbiol. Lett., 4, 133, 1978. 174. Liu, Y. et al. Confirmative electric DNA array-based test for food poisoning Bacillus cereus. J. Microbiol. Methods, 70, 55, 2007. 175. Ngamwongsatit, P. et al. WST-1-based cell cytotoxicity assay as a substitute for MTT-based assay for rapid detection of toxigenic Bacillus species using CHO cell line. J. Microbiol. Methods, 73, 211, 2008. 176. Magarvey, N.A., Ehling-Schulz, M. and Walsh, C.T. Characterization of the cereulide NRPS α-hydroxy acid specifying modules: activation of α-keto acids and chiral reduction on the assembly line. J. Am. Chem. Soc., 128, 10698, 2006. 177. Bennett, R.W. Staphylococcal enterotoxin and its rapid identification in foods by enzyme-linked immunosorbent assaybased methodology. J. Food Prot., 68, 1264, 2005.

11 Clostridium

Annamari Heikinheimo, Miia Lindström University of Helsinki

Dongyou Liu

Mississippi State University

Hannu Korkeala

University of Helsinki

Contents 11.1 Introduction.................................................................................................................................................................... 145 11.1.1 Classification..................................................................................................................................................... 145 11.1.2 Pathogenesis...................................................................................................................................................... 148 11.1.3 Diagnosis........................................................................................................................................................... 149 11.1.3.1 Foodborne C. perfringens................................................................................................................ 149 11.1.3.2 Foodborne C. botulinum.................................................................................................................. 150 11.2 Methods.......................................................................................................................................................................... 151 11.2.1 Sample Preparation........................................................................................................................................... 151 11.2.2 Detection Procedures........................................................................................................................................ 152 11.2.2.1 Multiplex PCR for C. perfringens Toxin Genes.............................................................................. 152 11.2.2.2 Multiplex PCR for C. botulinum Toxin Genes................................................................................ 152 11.2.2.3 PCR Assay to Distinguish Between Proteolytic (Mainly Group I) and Nonproteolytic (Mainly Group II) C. botulinum...................................................................................................... 152 11.3 Conclusion...................................................................................................................................................................... 153 References.................................................................................................................................................................................. 153

11.1  Introduction 11.1.1 Classification The genus Clostridium comprises a highly heterogeneous group of Gram-positive, endospore-producing, rod-shaped anaerobes that is ubiquitously distributed in the environment (e.g., soil, sewage, and marine sediments) and exists in the gastrointestinal tract of humans and domestic and feral animals. As obligate anaerobes, clostridia ferment by pathways that generate organic solvents, such as butyric acid, acetic acid, butanol, and acetone, as well as large amounts of gas (CO2 and H2) during fermentation of sugars. In addition, clostridia generate a variety of malodorous compounds during the fermentation of amino acids and fatty acids. The capability of clostridia to degrade large biological molecules (e.g., proteins, lipids, collagen, and cellulose) in the environment into fermentable components is undoubtedly assisted by their production of a wide range of extracellular enzymes. Thus, besides being useful for industrial production of alcohols and commercial solvents, clostridia play an important role in nature in biodegradation and the carbon cycle. Furthermore, a few species (e.g., C. butyricum and C. pasteurianum) are

capable of fixing nitrogen. Under stressful conditions, such as lack of nutrients or unfavorable temperatures, clostridia tend to produce spores that facilitate their persistence in the environment under extreme conditions. Although most Clostridium species are saprophytes, a small number (primarily C. perfringens, C. botulinum, C. tetani, and C. difficile) are pathogens to man and animals. These pathogenic species often secrete an array of invasins, exotoxins, and other extracellular enzymes (e.g., proteases, lipases, collagenase, and hyaluronidase). Various toxins are responsible for such human diseases as tetanus, botulism, and gas gangrene. C. perfringens (formerly called C. welchii) is a known toxin producer. The genome of C. perfringens strain 13 is 3,031,430 bp long, with 2,660 open reading frames and a low overall G + C content of 28.6%.1 Comparison of C. perfringens strain 13, gas gangrene isolate ATCC 13124T, and the enterotoxinproducing food poisoning strain SM101 has revealed considerable genomic diversity, with >300 unique genomic islands.2 Of the over 15 toxins produced by C. perfringens, alpha (CPA), beta (CPB), epsilon (ETX), and iota (ITX) toxins are the four major toxins present in five different toxinotypes (A–E) of the bacterium (Table 11.1).3 In addition, C. perfringens isolates 145

146

Molecular Detection of Foodborne Pathogens

Table 11.1 C. perfringens Toxinotypes and Their Major Toxins Toxinotype A B C D E

Alpha (CPA)

Beta (CPB)

Epsilon (ETX)

+ +

+

+ + + + +

may produce other toxins such as Clostridium perfringens enterotoxin (CPE) and Clostridium perfringens beta2 toxin (CPB2).4,5 These toxins have been also targeted for PCR identification (Table 11.2). CPE causes C. perfringens type A food poisoning as well as antibiotic-associated diarrhea (AAD) and sporadic diarrheas in humans.5 This toxin is also responsible for enteric diseases and enterotoxemias in animals.6 C. botulinum is a diverse species. The taxonomic denominator for C. botulinum is the production of botulinum neurotoxin (BoNT). C. botulinum is subdivided into four distinct groups based on physiological and genetic criteria, with Group I (proteolytic C. botulinum) strains producing one or two toxins of type A, B, or F; Group II (nonproteolytic C. botulinum) strains producing toxins of type B, E, or F; Group III strains producing toxins of type C or D; and Group IV strains producing toxin of type G.7 For the Group IV organisms, the species name Clostridium argentinense has been adopted.8 The genome of C. botulinum Group I (proteolytic) strain Hall A (ATCC 3502) consists of a chromosome of 3,886,916 bp and a plasmid of 16,344 bp, with 3,650 and 19 predicted genes, respectively.9 Clostridium sporogenes is phenotypically and genotypically related to Group I C. botulinum. Nontoxigenic type E-like counterparts of Group II C. botulinum have also been identified.10 C. botulinum secretes BoNT, which causes the deadly botulism food poisoning. BoNTs can be separated into seven distinct serotypes (A–G) on the basis of the immunological characteristics of its botulinum toxin. Moreover, four immunologically or genetically distinct BoNT/A subtypes, five BoNT/B subtypes, and six BoNT/E subtypes have been identified.11,12 While C. botulinum Groups I and II tend to cause foodborne botulism, C. botulinum Group I is often responsible for infant and wound botulism. Interestingly, strains of C. butyricum and C. baratii have also been shown to produce type E and F neurotoxins, respectively. Being adapted to a saprophytic lifestyle in both soil and aquatic environments, C. botulinum relies on the toxins and extracellular enzymes it produces to kill prey species, to gain access to nutrient sources, and to degrade rotting or decayed tissues.9 The strains of Group I C. botulinum are proteolytic and capable of utilizing amino acids as an energy source. The minimum growth temperature of Group I C. botulinum is 13–16°C and the optimum 37–42°C.13 Under otherwise favorable conditions, growth is typically inhibited by a water activity of 0.94, corresponding to approximately 10% NaCl (w:v) in brine. Growth does not occur under a pH of 4.5. Group II C. botulinum is nonproteolytic and saccharolytic. The minimum

Iota (ITX)

+ +

growth temperature is 3°C.14,15 The inhibitory water activity is 0.97, corresponding to 5% NaCl in brine. The spores of Group I C. botulinum possess a very high heat resistance.16 The spores of Group II C. botulinum are less heat-resistant than those of Group I. As Group I and II strains differ in their epidemiologies, discrimination between the two groups should be done whenever an outbreak strain is isolated. C. botulinum spores are widely spread in the environment and may contaminate raw foods. The heat-resistant Group I spores can cause problems in canning and especially in home preservation of vegetables and meat. Group II spores pose a safety hazard in modern food processing. They are able to grow at refrigeration temperatures, and anaerobic packages provide them with favorable conditions for growth and toxin formation. This sets great challenges for the production of modified atmosphere packaged chilled foods.17 C. tetani is the culprit for tetanus (lockjaw) in both humans and animals. C. tetani can be distinguished into 11 groups on the basis of its reaction with flagellar antigens. When C. tetani spores enter the skin via a puncture wound (e.g., stepping on a rusty nail), they germinate and form C. tetani vegetative cells. An exotoxin (called tetanus toxin or tetanospasmin) released by C. tetani cells migrates along neural paths and blocks the release of neurotransmitters (glycine and gamma-amino butyric acid) from the spinal cord and brainstem that regulate muscle contraction. This leads to continuous muscle ­contraction, primarily in the neck and jaw muscles (lockjaw) and respiratory failure, with mortality rates ranging from 40 to 78%. Being one of the three most poisonous substances known to humans (the other two are BoNT and diphtheria toxin), tetanus toxin (tetanospasmin, name of the gene encoded on a plasmid) comprises up to 5–10% of the bacterial cell weight and is heat-labile (destroyed at 56oC in 5 min) and O2-labile. C. difficile belongs to the normal microbiota of the mammalian gastrointestinal tract. C. difficile proliferation and infection in the human colon often occur after use of broadspectrum antibiotics. Through release of two toxins (A and B), C. difficile destroys the intestinal lining and causes AAD, colitis, and pseudomembranous colitis. C. difficile toxin A is an enterotoxin that induces fluid accumulation in the bowel, and toxin B is a cytopathic toxin that is extremely lethal. Both toxins are highly unstable and tend to degrade at room temperature. Additionally, C. septicum causes fatal traumatic (woundderived) and nontraumatic (endogenous) myonecrosis (malignant edema, gangrene) as well as enteric infections (usually a necrotic enteritis) in humans and animals. C. septicum cells

147

Clostridium

Table 11.2 Multiplex Primers for Determination of C. perfringens Toxinotypes Gene

Product (bp)

Present in Toxinotype

cpa

TGCATGAGCTTCAATTAGGT TTAGTTTTGCAACCTGCTGT

400

A–E

cpb

GCGAATATGCTGAATCATCTA GCAGGAACATTAGTATATCTTC

196

B, C

etx

GCGGTGATATCCATCTATTC CCACTTACTTGTCCTACTAAC

655

B, D

iA

ACTACTCTCAGACAAGACAG CTTTCCTTCTATTACTATACG

446

E

cpe

GGAGATGGTTGGATATTAGG GGACCAGCAGTTGTAGATA AGATTTTAAATATGATCCTAACC CAATACCCTTCACCAAATACTC

233

A–E

567

A–E

cpb2

Primers (probe)(5′–3′)

cpa

GTTGATAGCGCAGGACATGTTAAG CATGTAGTCATCTGTTCCAGCATC

402

A–E

cpb

ACTATACAGACAGATCATTCAACC TTAGGAGCAGTTAGAACTACAGAC

236

B, C

etx

ACTGCAACTACTACTCATACTGTG CTGGTGCCTTAATAGAAAGACTCC

541

B, D

itx

GCGATGAAAAGCCTACACCACTAC GGTATATCCTCCACGCATATAGTC

317

E

cpa

AGTCTACGCTTGGGATGGAA TTTCCTGGGTTGTCCATTTC

900

A–E

cpb

TCCTTTCTTGAGGGAGGATAAA TGAACCTCCTATTTTGTATCCCA

611

B, C

etx

TGGGAACTTCGATACAAGCA TTAACTCATCTCCCATAACTGCAC

396

B, D

iap

AAACGCATTAAAGCTCACACC CTGCATAACCTGGAATGGCT

293

E

cpb2

CAAGCAATTGGGGGAGTTTA GCAGAATCAGGATTTTGACCA

200

A–E

cpe

GGGGAACCCTCAGTAGTTTCA ACCAGCTGGATTTGAGTTTAATG

506

A–E

cpa

AAGAACTAGTAGCTTACATATCAACTAGTGGTG TTTCCTGGGTTGTCCATTTCC (VIC-TTGGAATCAAAACAAAGGATGGAAAAACTCAAG-TAMRA)

124

A–E

cpb

TGGAGCGTGAAAGAAACTGTTATTA GGTATCAAAAGCTAGCCTGGAATAGA (FAM-CTTAATTGGAATGGTGCTAACTGGGTAGGACAA-TAMRA)

85

B, C

etx

TTTGATAAGGTTACTATAAATCCACAAGGA AGAGAGCTTTTCCAACATAAACATCTTC (Cy-5-TAATCCTAAAGTTGAATTAGATGGAGAACCA-BHQ-2)

121

B, D

iap

GCATTAAAGCTCACACCTATTCCA GAGATGTGAGAGTTAATCCAAATTCTTG (FAM-CTAACTTAATTGTATATAGAAGGTCTGGTCC)

85

E

cpe

AGCTGCTGCTACAGAAAGATTAAATTT TGAGTTAGAAGAACGCCAATCATATAA (FAM-ACTGATGCATTAAACTCAAATCCAGCT-TAMRA)

88

A–E

cpb2

TATTTCAAAGTTTACTGTAATTTTTATGTTTTCA CCATTACCTTTCTATAAGCGTCGATT (Cy-5-TGCACTTGCTTTCATTGGACTTATTGCTCC-BHQ-2)

127

A–E

Reference

38–40

41

42

43

148

produces alpha toxin (molecular weight 46.5 kDa), probably via a type II secretion pathway, which is both hemolytic and lethal with 2 × 107 hemolytic units per milligram and an LD50 of approximately 10 μg/kg body weight in experimental mice. Interestingly, the C. septicum alpha toxin appears to be unique, as it is not found in any other clostridial species, although its mechanistic features are remarkably similar to those of the cytolytic toxin, aerolysin, from Aeromonas hydrophila. In addition, C. sordellii has been implicated in the deaths of a small number of women after childbirth. C. spiroforme causes rabbit diarrhea, resulting in significant losses in commercial rabbit farming facilities.18 In this chapter, we focus on two human foodborne pathogenic Clostridium species (i.e., C. perfringens and C. botulinum). Molecular methods for their rapid detection and identification are presented.

11.1.2  Pathogenesis C. perfringens type A is reported to be one of the most common causes of food poisonings throughout the industrialized world. The symptoms are caused by CPE, and although all types of C. perfringens (A–E) may produce CPE, usually food poisonings are caused by C. perfringens type A strains.19 C. perfringens type C occasionally causes enteritis necroticans (pigbel) in humans. The symptoms of this rare but fatal food poisoning are produced by C. perfringens beta toxin. C. perfringens type A food poisoning outbreaks are usually reported in institutionalized settings and involve numerous victims. Cooked meat and poultry are considered the most common vehicles for this food poisoning. After surviving in foods during preparation, C. perfringens type A spores germinate and the cells rapidly replicate in the food during cool-down and storage. The temperature range of 10–54°C allows the growth of the organism. Upon ingestion, C. perfringens cells sporulate in the small intestine and produce large quantities of CPE. As a consequence, severe abdominal cramps and diarrhea occur within 8–12 h, usually with a spontaneous recovery within 24 h. Fatalities are rare but possible in elderly and immunocompromised persons. On the molecular level, the enteropathogenic effects of CPE are based on its pore-forming capability in eukaryotic cells. CPE is a single polypeptide of 35 kDa (containing 319 amino acids). By first binding via protein receptors belonging to the claudin family (proteins involved in tight junction structure and function), CPE forms a large complex of 155 kDa. The continuing presence of CPE in intoxicated cells leads to the formation of another large complex of 200 kDa together with a tight junction protein called occludin. These CPE-containing complexes cause changes in cellular membrane permeability, resulting in death of the cells via either the apoptotic or oncotic pathways due to calcium influx through CPE pores.20 Interestingly, most C. perfringens type A food poisoning strains carry the cpe gene in their chromosome and

Molecular Detection of Foodborne Pathogens

show increased resistance to heating, osmotic stress, and low ­temperatures, whereas C. perfringens type A strains ­causing AAD, sporadic diarrhea, and animal diarrheas harbor the cpe gene in a plasmid and possess only moderate resistance to the above-mentioned food processing stresses.21,22 Evidence indicates that different subpopulations of cpe genecarrying (cpe-positive) C. perfringens type A strains exist and that the food poisoning strains seem to be adapted to the kitchen ­environment. However, recent findings show that also plasmidborne cpe-positive C. perfringens type A strains ­commonly cause food poisonings.23 Healthy food handlers have been ­demonstrated to serve as rich reservoirs for cpepositive C. perfringens type A, indicating that humans should be considered a possible source of contamination for C. perfringens type A food poisoning.24 C. botulinum produces BoNT during growth. BoNT is a single polypeptide chain of around 150 kDa. After nicking with bacterial protease (or gastric proteases), it turns into a light chain of 50 kDa and a heavy chain of 100 kDa. Being specific for peripheral nerve endings at the point where a motor neuron stimulates a muscle, BoNT binds to the neuron and prevents the release of acetylcholine across the synaptic cleft. The heavy chain mediates binding to ­presynaptic receptors via its carboxy terminus and forms a channel through the membrane of the neuron via its amino terminus, facilitating entrance of the light chain. Inside a neuron, the toxin specifically cleaves SNARE complex proteins (soluble NSF-attachment protein receptors [NSF, N-ethylmaleimide-sensitive fusion protein]), synaptobrevin (VAMP),25 ­synaptosomal protein (SNAP25),26,27 and syntaxin,28 which are involved in neurotransmitter release from synaptic vesicles. This abolishes the ability of affected cells to release a neurotransmitter, thus paralyzing the motor system. In structural and functional terms, BoNT is similar to tetanus toxin, as both are zinc-dependent endopeptidases that cleave a set of proteins in the synaptic vesicles of neurons involved in excretion of neurotransmitters. However, while BoNT displays a preference for stimulatory motor neurons at a neuromuscular junction in the peripheral nervous system, causing weakness or flaccid paralysis, tetanus toxin (tetanospasmin) acts on inhibitory motor neurons of the central nervous system, causing rigidity and spastic paralysis. Clinically, botulism often shows one of the four following manifestations: (i) foodborne botulism, which is caused by the ingestion of foods containing BoNT; (ii) infant botulism, which occurs in infants under one year of age after ingestion of C. botulinum spores that then colonize the intestinal tract and produce BoNT; (iii) wound botulism, which is caused by infection of C. botulinum in a wound, with BoNT being produced and spread to the body via the bloodstream; and (iv) intestinal colonization of C. botulinum in adults, which resembles infant botulism in its etiology. The paralysis typically starts in ocular and facial nerves, then proceeds to the throat, chest, and extremities, potentially leading to respiratory difficulty and death by asphyxia.

Clostridium

11.1.3  Diagnosis 11.1.3.1  Foodborne C. perfringens (i) Phenotypic identification: Being able to produce spores, the genus Clostridium is distinct among other anaerobic Gram-positive bacterial genera. However, this feature is often not discernible in uncultured specimens. Therefore, in vitro culture under strict anaerobic conditions followed by other phenotypic procedures is traditionally undertaken to identify Clostridium to species level. On blood agar plates, C. perfringens colonies are flat, rough, and translucent, with regular or irregular margins, an inner zone of hemolysis (resulting from the activity of perfringolysin O), and an outer zone of less complete clearing (caused by C. perfringens alpha toxin). Occasionally strains without inner zone hemolysis are seen. On Nagler agar containing 5–10% egg yolk, C. perfringens generates a characteristic white precipitate as a result of its alpha toxin (with lecithinase activity) interacting with the lipids in egg yolk. Several solid media have been developed for cultivation and isolation of C. perfringens from different sample materials. These include neomycin blood agar, sulfite polymyxin sulfadiazine (SPS) agar, tryptone sulfite neomycin (TSN) agar, Shahidi Ferguson perfringens (SFP) agar, oleandomycin polymyxin sulfadiazine perfringens (OPSP) agar, and tryptose sulfite cycloserine (TSC) agar with or without egg yolk.29–33 Isolation of typical colonies from media is followed by confirmation of C. perfringens colonies with different tests to reveal phenotypic characteristics. Observation of nitrate reduction, gelatine liquefaction, lactose fermentation, or lack of motility in suspected C. perfringens isolates is usually sufficient to distinguish C. perfringens from other organisms.34 Identification of C. perfringens may be also carried out using biochemical test kits such as API.33 Several international method standards are available for confirmation of C. perfringens in different samples, including methods published by the Association of Official Analytical Chemists (AOAC), the International Standardization Organization (ISO), and the Nordic Committee on Food Analysis (NCFA). Notably, not all C. perfringens isolates present in food or feces produce CPE and are thus capable of causing food poisonings. Therefore, detection of CPE production (or the presence of the cpe gene) in the suspected C. perfringens isolate is warranted when confirming the causative agent of the food poisoning. When detecting CPE production, a widely used technique is the enzyme-linked immunosorbent assay (ELISA).35,36 In addition, a kit based on reverse passive latex agglutination (RPLA) provides a straightforward method for detection of CPE in clinical samples.35,37 (ii) Molecular identification: Assays targeting C. perfringens ribosomal RNA (rRNA) and major toxin genes have been designed and applied in recent decades for improved detection and identification of this bacterium. Being a rapid, highly sensitive, and specific technique, PCR has become the method of choice for molecular confirmation of C. perfringens species and toxinotype

149

identity. Many PCR protocols focusing on individual C. perfringens toxin genes (cpa, cpb, etx, iap, cpe, and cpb2) have been reported. Further refinement has led to the development of multiplex PCR assays that enable simultaneous efficient detection of several C. perfringens toxin genes. For example, Meer and Songer38 described a multiplex PCR test that incorporates primers derived from C. perfringens cpa, cpb, cpe, etx, and iA genes for concurrent amplification of multiple target sequences (Table 11.2). Heikinheimo and Korkeala39 subsequently modified the assay. The subsequent inclusion of primers from C. perfringens cpb2 gene44 further enhanced the utility of this multiplex PCR for laboratory detection of the major toxinotypes of this bacterium.40 Yoo et al.41 also devised a multiplex PCR for C. perfringens major toxin genes cpa, cpb, etx, and itx, which is clearly superior to conventional seroneutralization with mice or guinea pigs. Using another independently developed multiple PCR targeting the alpha, beta, epsilon, and iota toxin genes, Garmory et al.44 effectively examined C. perfringens isolates from a variety of animals, including foals, piglets, and lambs. Augustynowicz et al.45 reported the use of primers for four different genes encoding the alpha, beta, epsilon, and iota toxins in a multiplex PCR for determination of genotype profile of C. perfringens isolates from Poland. The assay showed a sensitivity of 200 fg of DNA extracted from pure culture and a concordance of 94% between toxin phenotype and genotype. In another study, Baums et al.42 developed a reliable species-specific multiplex PCR for the detection of the cpa, cpb, cpb2, cpe, etx, and iap genes of C. perfringens isolates in a single reaction not requiring DNA purification (Table 11.2). With the goal of further speeding up the diagnostic process, Albini et al.43 reported the establishment and validation of three real-time fluorogenic (TaqMan) multiplex PCRs for rapid and precise identification of C. perfringens alpha, beta, beta2, epsilon, entero, and iotatoxin genes. This was achieved by using a combination of probe dyes (FAM/TAMRA, Cy-5/ BHQ-2, and VIC/TAMRA) and chromosome-borne alpha toxin as an internal positive control (Table 11.2). Besides showing total agreement with conventional PCR in identifying C. perfringens toxin genotypes, the newly developed multiplex PCRs provide the option of quantitating the copy numbers of plasmidborne toxin genes in relation to the chromosomally located alpha toxin gene. Similarly, Gurjar et al.46 described a real-time multiplex PCR utilizing a dual-labeled fluorescence hybridization probe (TaqMan®) for detection of C. perfringens toxin genes alpha (cpa), beta (cpb), iota (ia), epsilon (etx), beta2 (cpb2), and enterotoxin (cpe). This assay displayed a minimum detection limit of 5–70 pg of DNA depending on the toxin gene targets. In addition to PCR assays, other methods, such as DNA microarray, have also been reported for detecting cpe and other toxin genes.47,48 Colony hybridization has been used to detect cpe in raw beef.49 The authors developed a hydrophobic grid membrane filter colony hybridization method for the enumeration and isolation of cpe-positive C. perfringens

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directly from feces by detecting the cpe gene in bacterial colonies with a cpe-specific probe.50 (iii) Genetic typing: Typing is the assessment of the relatedness of a group of bacterial isolates to determine whether or not they originate from a common source. The ability to track C. perfringens type A strains involved in food poisoning outbreaks is vital for the control and prevention of this common illness. An early epidemiological tool in the investigation of C. perfringens type A food poisoning outbreaks was serotyping.51,52 This method is based on phenotypic characteristics of the organism, whereas current techniques rely to a great extent on genetic typing of the organism. A variety of nonamplified and amplified genetic typing techniques have been developed and applied. The nonamplified procedures include pulsedfield gel electrophoresis (PFGE), ribotyping, and plasmid profiling, whereas the amplified procedures include PCR, variable number tandem repeat (VNTR), and amplified fragment length polymorphism (AFLP). Nonamplified genetic typing methods. PFGE relies on gel electrophoresis to separate genomic C. perfringens DNA that has been previously digested with rare-cutting restriction enzymes, and generates a distinctive fingerprint pattern (consisting of five to 15 fragments in the range of 10–1,000 kb) for each strain. In assessing the clonality of 71 C. perfringens strains from 36 separate food poisoning outbreaks, Ridell et al.53 used SmaI and ApaI as restriction endonucleases. After evaluation of six restriction endonucleases (SmaI, ApaI, FspI, MluI, KspI, and XbaI), Maslanka et al.54 also chose SmaI, as this enzyme produced 11–13 well-distributed bands of 40–1,100 kb for effective discrimination of C. perfringens using PFGE. The authors obtained 17 distinct patterns among 62 isolates from seven outbreak investigations or control strains and noted that epidemiologically unrelated isolates had unique patterns, while isolates from different individuals within the same outbreak had similar, if not identical, patterns. Lukinmaa et al.55 reported the use of SmaI and ApaI in PFGE for typing 47 C. perfringens isolates from nine foodborne outbreaks of disease isolated during 1984–1999. Like many other bacterial species, C. perfringens harbors plasmids that can be targeted for genetic typing in a procedure called plasmid profiling.56–58 By isolating plasmid DNA from individual C. perfringens isolates and subjecting them to restriction enzyme digestion followed by electrophoresis, the plasmid profile (and genotype) of individual strains can be ascertained. For example, Eisgruber et al.57 employed plasmid profiling to show that the origin of three disease outbreaks could be clearly confirmed due to identical plasmid profiles in all C. perfringens isolates from food and clinical samples. Schalch et al.58 comparatively assessed plasmid profiling, ribotyping, and PFGE for typing 155 C. perfringens isolates from ten food poisoning outbreaks (with 34 food and fecal isolates) and 121 meat and fish pastes. They noted that although all three methods are suitable for classifying C. perfringens isolates below the species level, ribotyping, and PFGE generate patterns that can be interpreted more easily than plasmid profiling.

Molecular Detection of Foodborne Pathogens

Being a derivative of restriction fragment length polymorphism (RFLP), ribotyping uses a ribosomal DNA (rDNA) probe to detect the restriction fragment patterns of C. perfringens chromosomal DNA digested with appropriate restriction enzymes, providing much simpler and more consistent outcomes. Schalch et al.59 detected 12 ribopatterns with EcoRIdigested C. perfringens DNA from 34 isolates from 10 food poisoning cases and outbreaks over a seven-year period and found that all of the ribotypes of each food and stool isolate were identical in eight food poisoning cases and outbreaks. Amplified genetic typing methods. A key feature of amplified genetic typing methods is that C. perfringens­specific gene(s) of interest are amplified by PCR and analyzed directly by gel electrophoresis (e.g., VNTR) or subjected to other treatments before examination (e.g., AFLP). VNTRs, also known as microsatellites, are a class of short DNA sequence motifs that are tandemly repeated at certain specific locations, which play vital roles in both transcriptional and translational control of gene expression in bacteria and other organisms. Sawires and Songer60 characterized five VNTR loci in 112 C. perfringens isolates and demonstrated its value for rapid, easy, and cost-effective typing of this bacterium. In addition, this technique provides insight into the evolution of C. perfringens virulence.61 Chalmer et al.62 also employed VNTR in combination with capillary electrophoresis for precise and high-throughput typing of C. perfringens isolates. AFLP employs adaptors to restriction enzyme-digested C. perfringens DNA for PCR amplification and electrophoretic detection. McLauchlin et al.63 examined 35 C. perfringens clinical and food isolates from seven outbreaks and showed that AFLP is highly reproducible and provides a rapid, sensitive, and reproducible method for the genotyping of C. perfringens. Keto-Timonen et al.64 applied AFLP to differentiate C. perfringens strains from each other. The advantage of the AFLP method is that it overcomes the problem of extracellular DNAse production, which is often a problem when using, for example, PFGE for typing C. perfringens isolates. PCR genotyping assays are also available to establish whether cpe-positive C. perfringens type A isolates carry chromosomal or plasmidborne cpe. These methods detect different insertion sequence elements attached to cpe.65,66 11.1.3.2  Foodborne C. botulinum (i) Phenotypic identification: The complexity of the diagnostics of C. botulinum arises from the lack of validation of sensitive and specific in vitro methods for the detection of C. botulinum. The only standard method for detection of the neurotoxin is the mouse bioassay, which leads to ethical concerns regarding use of laboratory animals.67 Standard detection and isolation of C. botulinum is based on culture in a liquid medium and subsequent detection of BoNT in the culture supernatant by the mouse bioassay. Culture of C. botulinum requires strict anaerobic conditions. All culture media must be deoxygenated by heating in a boiling water bath or by a continuous flow of an anaerobic gas mixture, and the

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use of anaerobic jars and anaerobic workstations is required in a C. botulinum laboratory. Due to the extreme potency of BoNT, rigorous precautionary measures must be implemented to protect laboratory workers dealing with C. botulinum. Several sample pre-preparation steps can be utilized to enhance the isolation of C. botulinum from clinical samples (e.g., serum and feces). These include: (i) pretreatment with ethanol to eliminate vegetative bacteria and recover spores; (ii) heating to eliminate nonsporeforming bacteria (i.e., 80°C for 10 min for Group I spores or 60°C for 10–20 min for Group II spores); and (iii) inclusion of lysozyme (5 μg/ml) or other heat-resistant lytic enzymes in the culture medium to facilitate germination of heat-stressed spores.67 Chopped-meat-glucose-starch medium; cooked-meat medium; broths containing various combinations of tryptone, peptone, glucose, yeast extract, and trypsin (e.g., tryptonepeptone-glucose-yeast extract medium); reinforced clostridial medium; and fastidious anaerobe broth are widely used liquid media in cultivation of C. botulinum.68–73 Blood agar and egg yolk agar (EYA)74 are the most common unselective plating media, with EYA enabling the lipase reaction typical of C. botulinum. Group I strains grow optimally at 35–40°C, whereas Group II strains favor lower temperatures of 25–30°C.75 If both groups are being sought in a single sample, a compromise of 30°C has been proposed.75 However, investigating replicate samples at 26–30°C and at 35–37°C is preferable to ensure optimal growth for both groups. The optimal incubation time varies with the sample material and the detection method selected to identify the presence of C. botulinum in the culture tube. (ii) Molecular identification: As with C. perfringens, numerous PCR-based assays targeting the BoNT gene bot have been developed for the detection of C. botulinum. A multiplex PCR assay for C. botulinum botA, botB, botE, and botF genes has shown potential for rapid identification of C. botulinum toxinotypes of human botulism caused by Groups I and II76 (Table 11.3). However, the use of PCR assays targeting the neurotoxin genes does not provide details on the physiological group of C. botulinum isolates. In an attempt to address this issue, Dahlsten et al.77 designed a PCR assay targeting fldB, which is involved in phenylalanine metabolism in proteolytic clostridia, Table 11.3 Primers for C. botulinum Types A, B, E, and F Type A B E F

Primer (5′–3′)

Product Size (bp)

AGC TAC GGA GGC AGC TAT GTT CGT ATT TGG AAA GCT GAA AAG G CAG GAG AAG TGG AGC GAA AA CTT GCG CCT TTG TTT TCT TG CCA AGA TTT TCA TCC GCC TA GCT ATT GAT CCA AAA CGG TGA CGG CTT CAT TAG AGA ACG GA TAA CTC CCC TAG CCC CGT AT

782

Table 11.4 Primers for Differentiation between Group I and Group II C. botulinum Strains Gene fldB 16S rrn

Primer (5′–3′)

Product Size (bp)

GGGGAGAAAAAGTTGGTTGG CGAATCCACTAAATGGTGAAGG CGACCTGAGAGGGTGATCG GAGCTGACGACAACCATGC

552 761

to distinguish between proteolytic and nonproteolytic C. ­botulinum. The assay involves an internal amplification control targeted to the conserved region of 16S rrn in Group I and II C. botulinum (Table 11.4). This assay correctly identified all 36 Group I and 24 Group II C. botulinum strains, showing a 100% exclusivity and inclusivity. (iii) Genetic typing: Genetic typing of C. botulinum has been performed with various nonamplified and amplified procedures, including PFGE, ribotyping, AFLP, repetitive element sequence-based PCR (Rep-PCR), and VNTR. Lin and Johnson78 described the PFGE method using restriction enzymes MluI, RsrII, and SmaI for typing Group I A toxin-producing C. botulinum strains with encouraging results. Subsequently, Nevas et al.79 showed that use of SacII alone or in combination with SmaI and XhoI provides good discrimination among C. botulinum Group I strains. Others80,81 have demonstrated that restriction enzymes XhoI and SmaI can be employed in PFGE for genotyping of Group II C. botulinum strains. Application of ribotyping of EcoRI-digested C. botulinum DNA also permitted differentiation of C. botulinum strains belonging to Groups I and II.82 Keto-Timonen et al.83 showed that AFLP involving restriction enzymes HindIII and HpyCH4IV offers an excellent typing tool for Group I and II C. botulinum. Repetitive extragenic palindromes (REP) are randomly dispersed, repetitive sequence elements of 35–40 bp with an inverted repeat that exist in a wide range of bacterial genomes and are useful as genotyping targets. Hyytiä et al.84 reported a Rep-PCR protocol for determination of C. botulinum Group II type B and E toxin-producing strains to the strain level, and of C. botulinum Group II type F toxin-producing strains and all Group I strains to the toxinotype level. MacDonald et al.85 identified 10 VNTR regions in C. botulinum strain ATCC 3502 (subtype A1) for characterization of C. botulinum type A strains, resulting in the identification of 38 different genotypes among 59 strains. These markers have proven useful for discrimination among C. botulinum subtype A1 strains in the epidemiological investigations of botulism outbreaks.

205 389 543

11.2  Methods 11.2.1 Sample Preparation DNA extraction. One colony (C. perfringens or C. botulinum) is inoculated into 5 ml of cooked meat broth and

152

incubated at 26–30°C (Group II C. botulinum) or 37°C (C. perfringens and Group I C. botulinum) overnight (14 h). Cells are harvested by centrifugation (1,500 × g) at 4°C, and the pellet is resuspended in 600 μl of TE (10 mM Tris–HCl (pH 7.5), 1 mM EDTA (pH 8.0) containing 164 μg of RNase/ ml (Sigma), 8.3 mg of lysozyme/ml (Sigma), and 167 IU of mutanolysin/ml (Sigma) and incubated at 37°C for 30 min. Then, 9.5 mM EDTA (pH 8.0), 0.24 M NaCl, 49 μg of proteinase K/ml (Finnzymes, Espoo, Finland), and 0.8% (v/v) sodium dodecyl sulfate are added, and the mixture is incubated with gentle shaking at 60°C for 1 h to lyze the cells. Lysis is indicated by an increase in viscosity and a change in the suspension from turbid to opalescent. Subsequent phenol– chloroform–isoamyl alcohol (25:24:1, v/v) and chloroform– isoamyl alcohol (24:1, v/v) extractions are performed, and the DNA is precipitated with ethanol (95%, v/v), washed once with 70% ethanol, air-dried, and resuspended in 100 μl of 1 × TE (10 mM Tris–Cl pH 8.0, 1 mM EDTA pH 8.0) and then quantified with a spectrophotometer. For PCR analyses, the DNA is diluted in 1 × TE to a final concentration of 10 ng/μl. Alternatively, if C. perfringens and C. botulinum DNA are intended for PCR only, they can be prepared by using a boiling method. Namely, one to two colonies are suspended in 100 μl of water, heated at 95°C for 5 min, and centrifuged at 13,000 × g for 5 min, after which 1–5 μl is added to the PCR mixture (μl total volume).

11.2.2  Detection Procedures 11.2.2.1  Multiplex PCR for C. perfringens Toxin Genes Principle: All C. perfringens types (A–E) produce alpha toxin (encoded by the cpa gene). In addition, type B produces beta (encoded by the cpb gene) and epsilon (encoded by the etx gene) toxins, type C produces beta toxin, type D produces epsilon toxin, and type E produces iota toxin (encoded by the iA gene). Below, we present a multiplex PCR targeting C. perfringens cpa, cpb, etx, iA, and cpe genes, which was originally reported by Meer and Songer38 and recently updated by Heikinheimo and Korkeala.39 This assay provides a useful means of identifying C. perfringens toxigenotypes. Procedure: (1) Prepare the PCR mixture (50 µl) containing 10 μl of template, primers (0.34 mM of each cpe primer, 0.36 mM of each cpb primer, 0.46 mM of each etx primer, 0.5 mM of each cpa primer, and 0.52 mM of each iA primer), 1 × PCR buffer (10 mM Tris-HCl pH 8.3, 50 mM KCl, 1.5 mM MgCl2, and 0.001% gelatin), 0.1 mM of each dNTP, 5 U Taq DNA polymerase (Fisher Scientific), and distilled water (to 50 µl) (see Table 11.2).38,39 (2) Run the following cycling program in a PCR machine: one cycle of 94ºC for 2 min; 35 cycles of 94ºC for 1 min, 53ºC for 1 min, and 72ºC for 1 min; and one cycle of 72ºC for 10 min.

Molecular Detection of Foodborne Pathogens

(3) Separate the PCR products on a 1.5% agarose gel containing 0.5 μg/ml ethidium bromide and visualize under a UV light box and photograph. Note: the expected products are for C. perfringens type A 400 bp; type B 400, 196, and 655 bp; type C 400 and 196 bp; type D 400 and 655 bp; and type E 400 and 446 bp. C. perfringens cpe-positive strains show an additional 233-bp band. 11.2.2.2  Multiplex PCR for C. botulinum Toxin Genes Principle: C. botulinum causing human botulism belongs to two groups: (i) proteolytic Group I producing toxin A, B, or F or (ii) nonproteolytic Group II producing toxin B, E, or F. C. butyricum and C. baratii, which also produce type E and F toxins, respectively, can be distinguished from C. botulinum by biochemical methods. We present below a multiplex PCR assay for the simultaneous detection of type A1, B, E, and F neurotoxin genes76 (Table 11.3). Procedure: (1) Prepare the PCR mixture (50 μl) containing 0.3 μM concentrations of each primer (A, B, E, and F) (Table 11.3), 220 nM concentrations of each deoxynucleotide triphosphate (dATP, dCTP, dGTP, and dTTP; dNTP), 32 mM Tris-HCl (pH 8.4), 80 mM KCl, 4.8 mM MgCl, and 2 U of DNA polymerase (DynaZyme; Finnzymes), 1 μl of DNA template (see Section 11.2.1), and distilled water (to 50 μl). (2) Run the following cycling program in a PCR thermocycler: 28 cycles of 95ºC for 30 s, 60ºC for 25 s, and 72ºC for 1 min 25 s; and 1 cycle of 72ºC for 3 min. (3) Separate the PCR products on a 2% agarose gel containing 0.5 μg/ml ethidium bromide along with a DNA molecular weight marker (e.g., DNA molecular weight marker VI, Roche Diagnostics) and visualize under a UV light box and photograph. Note: a PCR product of 782 bp is observed with C. botulinum subtype A1 alone, 205 bp for type B alone, 389 bp for type E alone, and 543 bp for type F alone. Other bacterial species, including C. botulinum subtypes A2–A4, C. sporogenes, and the nontoxigenic nonproteolytic C. botulinum-like organisms, do not yield a PCR product. Sensitivity of the PCR for types A, E, and F is 102 cells and for type B ten cells per reaction mixture. With a two-step enrichment, the detection limit in food and fecal samples varied from 102 spore/g for types A, B, and F to 101 spore/g of sample material for type E.76 11.2.2.3 PCR Assay to Distinguish Between Proteolytic (Mainly Group I) and Nonproteolytic (Mainly Group II) C. botulinum Principle: Dahlsten et al.77 developed a multiplex PCR with primers targeted to the fldB gene (associated with

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Clostridium

phenylalanine metabolism in proteolytic clostridia) for specific recognition of Group I C. botulinum, and to a conserved region of 16S rrn (present in both Group I and Group II C. botulinum) (Table 11.4). The 16S rrn primers act as an internal control for both Group I and II C. botulinum.



Procedure: (1) Prepare the PCR mixture (50 μl) containing 1 × PCR buffer (10 mM Tris–HCl, 1.5 mM MgCl2, 50 mM KCl, 0.1% Triton X-100 (DyNAzyme Reaction Buffer, Finnzymes), 200 μM of each dNTP, 0.5 μM of each fldB primer, 0.1 μM of each 16S rnn primer, 2 U Taq DNA polymerase (Finnzymes), 5 μl of DNA, and distilled water (to 50 μl). (2) Run the following cycling program in a PCR machine: one cycle of 95ºC for 2 min; 28 cycles of 95ºC for 30 s, 53ºC for 30 s, and 72ºC for 1 min; and 1 cycle of 72ºC for 5 min. (3) Separate the PCR products on a 2% agarose gel containing 0.5 μg/ml ethidium bromide at 90 V for 1 h 45 min along with a DNA molecular weight marker (e.g., DNA molecular weight marker VI, Roche Diagnostics) and visualize under a UV light box and photograph. Note: the expected products are for C. botulinum Group I 552 and 752 bp; and for Group II 752 bp.77 The assay is intended for rapid typing (Group I vs. Group II) of pure isolates previously identified as C. botulinum. The assay is not suitable for analysis of mixed cultures.









11.3  Conclusion Besides causing serious nonfoodborne diseases, C. perfringens and C. botulinum contribute to an increasing number of food-related outbreaks worldwide. The ability to correctly identify these bacteria and differentiate between their toxigenotypes is vital to the control and prevention of diseases arising from them. While the conventional culture techniques are useful for isolation and identification of these bacteria, these methods are often time-consuming and sometimes variable. The application of molecular procedures has greatly improved laboratory diagnostics of these important bacterial pathogens. There is no doubt that continuing refinement in the sample processing procedures and automation in the target amplification and detection will make molecular identification of C. perfringens and C. botulinum and their toxinotypes an indispensable tool in food and clinical testing laboratories, and further strengthen our capacity in the tracking and prevention of foodborne illnesses resulting from C. perfringens and C. botulinum infection and contamination.

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1. Shimizu, T. et al. Complete genome sequence of Clostridium perfringens, an anaerobic flesh-eater. Proc. Natl. Acad. Sci. USA, 99, 996, 2002.







2. Myers, G.S. et al. Skewed genomic variability in strains of the toxigenic bacterial pathogen, Clostridium perfringens. Genome Res., 16, 1031, 2006. 3. Hatheway, C.L. Toxigenic clostridia. Clin. Microbiol. Rev., 3, 66, 1990. 4. Gibert, M., Jolivetrenaud, C. and Popoff, M.R. Beta2 toxin, a novel toxin produced by Clostridium perfringens. Gene, 203, 65, 1997. 5. Smedley, J.G. 3rd et al. The enteric toxins of Clostridium perfringens. Rev. Physiol. Biochem. Pharmacol., 152, 183, 2005. 6. Songer, J.G. and Uzal, F.A. Clostridial enteric infections in pigs. J. Vet. Diagn. Invest., 17, 528, 2005. 7. Peck, M.W. Clostridium botulinum and the safety of minimally heated, chilled foods: An emerging issue? J. Appl. Microbiol., 101, 556, 2006. 8. Suen, J.C. et al. Clostridium argentinense, sp. nov: A genetically homogenous group composed of all strains of Clostridium botulinum type G and some nontoxigenic strains previously identified as Clostridium subterminale or Clostridium hastiform. Int. J. Syst. Bacteriol., 38, 375, 1988. 9. Sebaihia, M. et al. Genome sequence of a proteolytic (Group I) Clostridium botulinum strain Hall A and comparative analysis of the clostridial genomes. Genome Res., 17, 1082, 2007. 10. Kautter, D.A. et al. Antagonistic effect on Clostridium botulinum type E by organisms resembling it. Appl. Microbiol., 14, 616, 1966. 11. Hill, K.K. et al. Genetic diversity among botulinum neurotoxinproducing clostridial strains. J. Bacteriol., 189, 818, 2007. 12. Chen, Y. et al. Sequencing the botulinum neurotoxin gene and related genes in Clostridium botulinum type E strains reveals orfx3 and a novel type E neurotoxin subtype. J. Bacteriol., 189, 8643, 2007. 13. Hinderink, K., Lindström, M. and Korkeala, H. Group I Clostridium botulinum strains show significant variation in growth at low and high temperature. J. Food Prot, 72(1) In press. 14. Eklund, M.W., Wieler, D.I. and Poysky, F.T. Outgrowth and toxin production of nonproteolytic type B Clostridium botulinum at 3.3 to 5.6°C. J. Bacteriol., 93, 1461, 1967. 15. Graham, A.F., Mason, D.R., and Peck, M.W. Effect of pH and NaCl on growth from spores of nonproteolytic Clostridium botulinum at chill temperature. Lett. Appl. Microbiol., 24, 95, 1997. 16. Stumbo, C.R., Purohit, K.S. and Ramakrishna, T.V. Thermal process lethality guide for low-acid foods in metal containers. J. Food Sci., 40, 1316, 1975. 17. Lindström, M., Kiviniemi, K. and Korkeala, H. Hazard and control of group II (non-proteolytic) Clostridium botulinum in modern food processing. Int. J. Food Microbiol., 108, 92, 2006. 18. Drigo, I. et al. Development of PCR protocols for specific identification of Clostridium spiroforme and detection of sas and sbs genes. Vet Microbiol., 2008. 19. McClane, B.A. Clostridium perfringens. In: Food microbiology: Fundamentals and frontiers, 2nd ed. p. 351. ASM Press, Washington DC, 2001. 20. Smedley, J.G. et al. Identification of a prepore large-complex stage in the mechanism of action of Clostridium perfringens enterotoxin. Infect. Immun., 75, 2381, 2007. 21. Li, J. and McClane, B.A. Further comparison of temperature effects on growth and survival of Clostridium perfringens type A isolates carrying a chromosomal or plasmid-borne enterotoxin gene. Appl. Environ. Microbiol., 7, 4561, 2006.

154 22. Li J. and McClane, B.A. Comparative effects of osmotic, sodium nitrite-induced, and pH-induced stress on growth and survival of Clostridium perfringens type A isolates carrying chromosomal or plasmid-borne enterotoxin genes. Appl. Environ. Microbiol., 12, 7620, 2006. 23. Lahti, P. et al. Clostridium perfringens type A strains carrying a plasmid-borne enterotoxin gene (genotype IS1151-cpe or IS1470-like-cpe) as a common cause of food poisoning. J. Clin. Microbiol., 46, 371, 2008. 24. Heikinheimo, A. et al. Humans as reservoir for enterotoxin gene-carrying Clostridium perfringens type A. Emerg. Infect. Dis., 12, 1724, 2006. 25. Schiavo, G. et al. Tetanus and botulinum-B neurotoxins block transmitter release by proteolytic cleavage of synaptobrevin. Nature, 359, 832, 1992. 26. Blasi, J. et al. Botulinum neurotoxin A selectively cleaves the synaptic protein SNAP-25. Nature, 365, 160, 1993. 27. Schiavo, G. et al. Botulinum neurotoxins serotypes A and E cleave SNAP-25 at distinct COOH-terminal peptide bonds. FEBS Lett., 335, 99, 1993. 28. Schiavo, G. et al. Botulinum neurotoxin type C cleaves a single Lys-Ala bond within the carboxyl-terminal region of syntaxins. J. Biol. Chem., 270, 10566, 1995. 29. Harmon, S.M., Kautter, D.A. and Peeler, J.T. Improved medium for enumeration of Clostridium perfringens. Appl. Microbiol., 22, 688, 1971. 30. Hauschild, A.H. and Hilsheimer, R. Enumeration of foodborne Clostridium perfringens in egg yolk-free tryptosesulfite-cycloserine agar. Appl. Microbiol., 27, 521, 1974. 31. Hauschild, A.H. and Hilsheimer, R. Evaluation and modifications of media for enumeration of Clostridium perfringens. Appl. Microbiol., 27, 78, 1974. 32. Hauschild, A.H., Hilsheimer, R. and Griffith, D.W. Enumeration of fecal Clostridium perfringens spores in egg yolk-free tryptose-sulfite-cycloserine agar. Appl. Microbiol., 27, 527, 1974. 33. Labbé, R. Clostridium perfringens. In: The microbiological safety and quality of food. Aspen Publishers, Gaithersburg, MD, 2000. 34. Cato, E., George, W.L. and Finegold, S. Genus Clostridium, Bergey’s manual of systematic bacteriology. Vol. 2. Williams and Wilkins, Baltimore, MD, USA, 1986. 35. Berry, P.R. et al. Evaluation of ELISA, RPLA, and Vero cell assays for detecting Clostridium perfringens enterotoxin in faecal specimens. J. Clin. Pathol., 41, 458, 1988. 36. Piyankarage, R.H. et al. Sandwich enzyme-linked immunosorbent assay by using monoclonal antibody for detection of Clostridium perfringens enterotoxin. J. Vet. Med. Sci., 61, 45, 1999. 37. Berry, P.R. and Stringer, M.F. Preliminary investigation on the use of latex agglutination for the detection of Clostridium perfringens enterotoxin. J. Appl. Bacteriol., 59, 14, 1985. 38. Meer, R. and Songer, G. Multiplex polymerase chain reaction assay for genotyping Clostridium perfringens. Am. J. Vet. Res., 58, 702, 1997. 39. Heikinheimo, A. and Korkeala, H. Multiplex PCR assay for toxinotyping Clostridium perfringens isolates obtained from Finnish broiler chickens. Lett. Appl. Microbiol., 40, 407, 2005. 40. Uzal, F.A. and Songer, J.G. Diagnosis of Clostridium perfringens intestinal infections in sheep and goats. J. Vet. Diagn Invest., 20, 253, 2008. 41. Yoo, H.S. et al. Molecular typing and epidemiological survey of prevalence of Clostridium perfringens types by multiplex PCR. J. Clin. Microbiol., 35, 228, 1997.

Molecular Detection of Foodborne Pathogens 42. Baums, C.G. et al. Diagnostic multiplex PCR for toxin genotyping of Clostridium perfringens isolates. Vet. Microbiol., 100, 11, 2004. 43. Albini, S. et al. Real-time multiplex PCR assays for reliable detection of Clostridium perfringens toxin genes in animal isolates. Vet. Microbiol., 127, 179, 2008. 44. Garmory, H.S. et al. Occurence of Clostridium perfringens β2-toxin amongst animals, determined using genotyping and subtyping PCR assays. Epidemiol. Infect., 124, 61, 2000. 45. Augustynowicz, E., Gzyl, A. and S´lusarczyk, J. Molecular epidemiology survey of toxinogenic Clostridium perfringens strain types by multiplex PCR. Scand. J. Infect. Dis., 32, 637, 2000. 46. Gurjar, A.A. et al. Real-time multiplex PCR assay for rapid detection and toxintyping of Clostridium perfringens toxin producing strains in feces of dairy cattle. Mol. Cell. Probes, 22, 90, 2008. 47. Al-Khaldi, S.F. et al. Genotyping of Clostridium perfringens toxins using multiple oligonucleotide microarray hybridization. Mol. Cell. Probes, 18, 359, 2004. 48. Al-Khaldi, S.F. et al. Identification and characterization of Clostridium perfringens using single target DNA microarray chip. Int. J. Food Microbiol., 91, 289, 2004. 49. Baez, L.A. and Juneja, V.K. Nonradioactive colony hybridization assay for detection and enumeration of enterotoxigenic Clostridium perfringens in raw beef. Appl. Environ. Microbiol., 61, 807, 1995. 50. Heikinheimo, A., Lindström, M. and Korkeala, H. Enumeration and isolation of cpe-positive Clostridium perfringens spores from feces. J. Clin. Microbiol., 42, 3992, 2004. 51. Hauschild, A.H.W. Criteria and procedures for implicating Clostridium perfringens in food-borne outbreaks. Can. J. Public Health, 66, 388, 1975. 52. Lund, B.M., Baird-Parker, T.C. and Gould, G.W. The microbiological safety and quality of food. Aspen Publishers, Gauthersburg, MD, 2000. 53. Ridell, J. et al. Prevalence of the enterotoxin gene and clonality of Clostridium perfringens strains associated with foodpoisoning outbreaks. J. Food Prot., 61, 240, 1998. 54. Maslanka, S.E. et al. Molecular subtyping of Clostridium perfringens by pulsed-field gel electrophoresis to facilitate foodborne-disease outbreak investigations. J. Clin. Microbiol., 37, 2209, 1999. 55. Lukinmaa, S., Takkunen, E. and Siitonen, A. Molecular epidemiology of Clostridium perfringens related to food-borne outbreaks of disease in Finland from 1984 to 1999. Appl. Environ. Microbiol., 68, 3744, 2002. 56. Mahony, D.E., Ahmed, R. and Jackson, S.G. Multiple typing techniques applied to a Clostridium perfringens food poisoning outbreak. J. Appl. Bacteriol., 72, 309, 1992. 57. Eisgruber, H., Wiedmann, M. and Stolle, A. Use of plasmid profiling as a typing method for epidemiologically related Clostridium perfringens isolates from food poisoning cases and outbreaks. Lett. Appl. Microbiol., 20, 290, 1995. 58. Schalch, B. et al. Molecular methods for the analysis of Clostridium perfringens relevant to food hygiene. FEMS Immunol. Med. Microbiol., 24, 281, 1999. 59. Schalch, B. et al. Ribotyping for strain characterization of Clostridium perfringens isolates from food poisoning cases and outbreaks. Appl. Environ. Microbiol., 63, 3992, 1997. 60. Sawires, Y.S. and Songer, J.G. Multiple-locus variable-number tandem repeat analysis for strain typing of Clostridium perfringens. Anaerobe, 11, 262, 2005.

Clostridium 61. Sawires Y.S. and Songer, J.G. Clostridium perfringens: Insight into virulence evolution and population structure. Anaerobe, 12, 23, 2006. 62. Chalmers, G. et al. Typing of Clostridium perfringens by multiple-locus variable number of tandem repeats analysis. Vet. Microbiol., 128, 126, 2008. 63. McLauchlin, J. et al. Amplified fragment length polymorphism (AFLP) analysis of Clostridium perfringens for epidemiological typing. Int. J. Food Microbiol., 56, 21, 2000. 64. Keto-Timonen, R. et al. Identification of Clostridium species and DNA fingerprinting of Clostridium perfringens by amplified fragment length polymorphism analysis. J. Clin. Microbiol., 44, 4057, 2006. 65. Wen, Q.Y., Miyamoto, K. and McClane, B.A. Development of a duplex PCR genotyping assay for distinguishing Clostridium perfringens type A isolates carrying chromosomal enterotoxin (cpe) genes from those carrying plasmid-borne enterotoxin (cpe) genes. J. Clin. Microbiol., 41, 1494, 2003. 66. Miyamoto, K., Wen, Q.Y. and McClane, B.A. Multiplex PCR genotyping assay that distinguishes between isolates of Clostridium perfringens type A carrying a chromosomal enterotoxin gene (cpe) locus, a plasmid cpe locus with an IS1470-like sequence, or a plasmid cpe locus with an IS1151 sequence. J. Clin. Microbiol., 42, 1552, 2004. 67. Lindström, M. and Korkeala, H. Laboratory diagnostics of botulism. Clin. Microbiol. Rev., 19, 298, 2006. 68. Robertson, M. Notes upon certain anaerobes isolated from wounds. J. Pathol. Bacteriol., 20, 327, 1916. 69. Gibbs, B.M. and Hirsch, A. Spore formation by Clostridium species in an artificial medium. J. Appl. Bacteriol., 19, 129, 1956. 70. Lilly, T. et al. An improved medium for detection of Clostridium botulinum type E. J. Milk Food Technol., 34, 492, 1971. 71. Quagliaro, D.A. An improved cooked meat medium for the detection of Clostridium botulinum. J. AOAC, 60, 563, 1977. 72. Centers for Disease Control and Prevention. Botulism in the United States (1899–1996). Handbook for epidemiologists, clinicians, and laboratory workers. U.S. Department of Health, Education, and Welfare, CDC, Atlanta, GA, 1998. 73. Saeed, E.M.A. Studies on isolation and identification of Clostridium botulinum investigating field samples specially from equine grass sickness cases. Doctoral thesis, University of Göttingen, Göttingen, Germany, 2005.

155 74. Hauschild, A.H.W. and Hilsheimer, R. Enumeration of Clostridium botulinum spores in meats by a pour-plate procedure. Can. J. Microbiol., 23, 829, 1977. 75. Smith, L.D.S. and Sugiyama, H. Botulism. The organism, its toxins, the disease. Charles C. Thomas, Springfield, IL, 1988. 76. Lindström, M. et al. Multiplex PCR assay for detection and identification of Clostridium botulinum types A, B, E, and F in food and faecal material. Appl. Environ. Microbiol., 67, 5694, 2001. 77. Dahlsten, E. et al. PCR assay for differentiating between Group I (proteolytic) and Group II (nonproteolytic) strains of Clostridium botulinum. Int. J. Food Microbiol., 124. 108, 2008. 78. Lin, W.J. and Johnson, E.A. Genome analysis of Clostridium botulinum type A by pulsed-field gel electrophoresis. Appl. Environ. Microbiol. 61, 4441, 1995. 79. Nevas, M. et al. Diversity of proteolytic Clostridium botulinum strains, determined by pulsed-field gel electrophoresis approach. Appl. Environ. Microbiol., 71, 1311, 2005. 80. Hielm, S. et al. Genomic analysis of Clostridium botulinum group II by pulsed-field gel electrophoresis. Appl. Environ. Microbiol., 64, 703, 1998. 81. Hyytiä, E. et al. Biodiversity of Clostridium botulinum type E strains isolated from fish and fishery products. Appl. Environ. Microbiol., 65, 2057, 1999. 82. Hielm, S. et al. Ribotyping as an identification tool for Clostridium botulinum strains causing human botulism. Int. J. Food Microbiol., 47, 121, 1999. 83. Keto-Timonen, R., Nevas, M. and Korkeala, H. Efficient DNA fingerprinting of Clostridium botulinum types A, B, E, and F by amplified fragment length polymorphism analysis. Appl. Environ. Microbiol., 71, 1148, 2005. 84. Hyytiä, E. et al. Characterisation of Clostridium botulinum groups I and II by randomly amplified polymorphic DNA analysis and repetitive element sequence-based PCR. Int. J. Food Microbiol., 48, 179, 1999. 85. MacDonald, T.E. et al. Differentiation of Clostridium botulinum serotype A strains by multiple-locus variable-number tandem-repeat analysis. Appl. Environ. Microbiol., 74, 875, 2008.

12 Enterococcus

Teresa Semedo-Lemsaddek, Rogério Tenreiro University of Lisbon

Paula Lopes Alves, Maria Teresa Barreto Crespo

Institute of Experimental and Technological Biology (IBET)

Contents 12.1 Introduction.................................................................................................................................................................... 157 12.1.1 Taxonomy and General Characteristics of Enterococci................................................................................... 157 12.1.2 Molecular Approaches for Identification and Detection of Enterococci.......................................................... 158 12.1.3 Why Detect and/or Quantify Enterococci?....................................................................................................... 168 12.2 Methods.......................................................................................................................................................................... 168 12.2.1 Sample Collection and Preparation.................................................................................................................. 168 12.2.1.1 Sample Homogenization.................................................................................................................. 168 12.2.1.2 Cell Isolation and Counting without Pre-enrichment...................................................................... 169 12.2.1.3 Pre-enrichment................................................................................................................................ 169 12.2.1.4 DNA Extraction for Conventional PCR........................................................................................... 169 12.2.1.5 DNA Extraction for Real-time PCR.................................................................................................170 12.2.1.6 DNA Extraction from a Fermented Cereal-based Food ................................................................. 172 12.2.2 Detection Procedures.........................................................................................................................................173 12.2.2.1 Genus- and Species-specific Multiplex PCR....................................................................................173 12.2.2.2 Data Analysis....................................................................................................................................174 12.3 Conclusions and Future Perspectives..............................................................................................................................174 Acknowledgments.......................................................................................................................................................................175 References...................................................................................................................................................................................175

12.1  Introduction 12.1.1 Taxonomy and General Characteristics of Enterococci The classification and nomenclature of the genus Enterococcus have attracted interest and constant discussion over the years, since the first proposal of the name “enterocoques” made in 1902 by Thiercelin.1 In 1970, Kalina2 proposed that Streptococcus faecalis and S. faecium should be renamed as Enterococcus faecalis and E. faecium, respectively. These proposals remained unrecognized officially and in 1980 the genus Enterococcus did not appear in the Approved List of Bacterial Names. In the 1980s, Schleifer and Kilpper-Bälz3 resurrected the genus Enterococcus after performing DNA–DNA and DNA–rRNA hybridizations. Other studies carried out in the same decade, including 16S rRNA sequencing,4 also demonstrated that enterococci form a separate genus. Analysis of these molecular data divided the streptococci sensu lato into three genera: (i) S. sensu stricto, comprising the majority of the known species; (ii) Enterococcus, for the enteric group; and (iii) Lactococcus, for the lactic streptococci.

Since then the genus Enterococcus has been accepted as valid and today it comprises 33 species. In 2007 Kohler5 performed an extensive review of the publications that lead to each of the species currently accepted and included in Taxonomic Outline of the Bacteria and Archaea (TOBA), based on a comprehensive taxonomy using 16S rRNA gene phylogeny.6 Members of the genus Enterococcus are Gram-positive ovoid cells that appear single, in pairs or short chains, catalase, and oxidase negative, facultative anaerobes, and homofermentative with lactic acid as the end product of glucose fermentation. The majority of the enterococci grow from 10 to 45ºC, with 6.5% NaCl and at pH 9.6, hydrolyze esculin in the presence of 40% bile salts and the G+C content ranges from 37 to 45%.7,8 However, not all enterococci exhibit these characteristics with several authors analyzing and reporting differences in physiological properties among different species and strains belonging to the same species.9–14 Enterococci encompass a large proportion of the autochthonous gastrointestinal microbiota of humans and animals and can also be found in soil, surface waters, plant material, vegetables and raw, and fermented food.14–18 157

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Enterococci are known to be hardy microorganisms able to resist to pasteurization temperatures and adapt to different substrates and growth conditions (low and high temperature, wide pH range and salinity). This high adaptability implies that they can be found either in food products manufactured from raw materials (e.g., milk or meat) or in heat-treated food products. Studies performed with cheese from different origins, namely France,19,20 Greece,21,22 Italy,23–26 Portugal,27,28 Slovenia,29 Spain,30 and Ireland31 demonstrated that enterococci are a relevant part of the indigenous microbiota and contribute to the development of sensory characteristics.24,32,33 These bacteria are also associated with fermented meat products,15,34,35 vegetables and olives,35–37 and plants in general.38,39 In meat products the most frequent species to be detected are E. faecalis and E. faecium and less frequently or occasionally found are the species E. durans, E. hirae, E. gallinarum, E. casseliflavus, and E. mundtii. In cheeses the frequency of appearance varies according to the type of milk and country of origin but the most frequently reported species is E. faecium, followed by E. faecalis, E. durans, E. hirae, and E. casseliflavus. Products from plant origin do not differ substantially on the enterococcal species found, namely E. faecium, E. faecalis, E. casseliflavus, and E. flavescens. Another benefit of enterococci in food is that many produce bacteriocins with a broad antimicrobial spectrum against pathogenic bacteria.24,40 However, although enterococci belong to the lactic acid bacteria, they are not “generally recognized as safe” (GRAS) organisms limiting their application as starters or probiotics.13,33,41 Although some desirable metabolic traits turn enterococci essential for the production of many food products, the occurrence of antimicrobial resistance and virulence factors, associated with their highly efficient ability to transfer genetic material, pose enterococci as an emerging threat as human pathogens associated with nosocomial and community acquired infections. Strains with virulence determinants and antimicrobial resistance have been isolated not only from medical wards, human or veterinarian, but also from environmental and food samples. 14,17,33,42–46 Among enterococcal species E. faecalis, and to a lesser extent E. faecium, appear as predominantly associated with human pathogenesis with special incidence of vancomycinresistant strains.47–50 Other enterococcal species such as E. avium, E. ­casseliflavus, E. durans, E. gallinarum, E. gilvus, E. hirae, E. malodoratus, E. mundtii, E. pallens, E. raffinosus, and E. solitarius have been reported as infrequent causes of human infection.32,51–56 Since the majority of the enterococcal infections are associated with E. faecalis and E. faecium, the accurate identification at the species level becomes important for surveillance of food products, hospitals, and other environments.

12.1.2 Molecular Approaches for Identification and Detection of Enterococci The presence of microorganisms in food is a natural and unavoidable occurrence. Cooking generally destroys

Molecular Detection of Foodborne Pathogens

most harmful bacteria, but undercooked foods, processed ­ready-to-eat foods and minimally processed foods can contain harmful bacteria that constitute serious health threats. Researchers are continuously searching for tools that are fast, accurate and ultrasensitive to detect the presence of these microorganisms. Conventional culture methods remain the most reliable and accurate techniques for foodborne pathogen detection. Traditionally, enterococcal identification relied on various cultivation procedures associated with the analysis of phenotypic profiles including Gram staining, colony morphology, motility, pigmentation, growth requirements and enzymatic, and/or metabolic activities. However, this culture-based identification is time-consuming and frequently limited by the variability of atypical strains, the lack of sufficient data on recently described species and the low reproducibility of some tests. Since Enterococcus is not a phylogenetically nor phenotypically coherent and homogeneous genus, manual or automated systems based on phenotypic characteristics not always allow the correct strain identification, therefore these problems assume great importance.8 Another disadvantage of the standard culture methods is their lack of ability to detect a substantial proportion of the bacterial population which is nonculturable. These viable but nonculturable (VBNC) microorganisms demonstrate metabolic activity,57,58 maintain their pathogenicity59,60 and their antibiotic resistance traits,61 can express,62 and exchange genes,63 and are able to reestablish division upon restoration of favorable conditions.64,65 It has recently been demonstrated that E. faecalis can enter the VBNC state,62 thus representing a risk for food safety and emphasizing the importance of molecular detection in substitution of standard culture methods. Consequently, the optimization of molecular methods for microbial analysis is becoming more important every day, with special relevance for rapid, easy to implement and reliable procedures that may be applied in routine laboratories to ensure the safety and quality of food. Nucleic acid based identification and detection methods have been developed for nearly all bacterial pathogens.66 Rapid and sensitive DNA-based assays which are applicable for the direct detection of enterococci in food, as well as in clinical or environmental samples, may improve the quickness and the accuracy of the detection of these bacteria. In spite of all the advantages associated with molecularbased methodologies, the detection of microorganisms in food products can come across several problems, such as PCR inhibition by food components, sample cross-contamination in sensitive PCR assays, detection of living as well as dead cells (this specific problem can be solved by using mRNA as amplification target instead of DNA in RT-PCR). Additionally, PCR may be too sensitive for some applications, detecting a microbe that is present at nonpathogenic levels. Although the above mentioned problems, and others, have to be considered when optimizing DNA-based methods, PCR amplification of species-specific targets enables precise identification of known microbial species with short reporting times. However, it is also important

Enterococcus

to highlight that positive amplification only demonstrates the presence of the appropriate target DNA sequences in the sample and does not imply that the target organism is viable. This problem can be overcome by the evaluation of cell viability (mRNA detection by RT-PCR) or visualization of living cells in mixed bacterial populations (in situ PCR/hybridization). Several molecular diagnostic methods based on the detection of bacterial nucleic acids have already been described for enterococci. The majority of these methodologies relies on or includes PCR due to the low amounts of DNA and/or cells needed, as well as the detection of a specific DNA sequence against a large background of prokaryotic and eukaryotic cells and organic material present in mixed samples.67 Under ideal conditions, with 100% efficiency and unlimited resources, a single copy of target gene can be amplified to approximately 3.4 × 1010 copies after 35 cycles of PCR. However, it becomes important to emphasize that the ability to design oligonucleotide primers is limited by our knowledge of a microorganism’s genome, as well as by the representativeness of publicly available sequence databases in terms of all variants of that microbial group. Distinct molecular approaches have already been described for enterococci including broad-range PCR,68 multiplex PCR,69–71 species-specific PCR,72–77 real-time PCR,66,78,79 direct detection of bacteria using fluorescent in situ hybridization (FISH) probes80–82 and amplification of 16S and 23S ribosomal gene sequences for hybridization to a microarray.83,84 Conventional PCR assays detect only the presence or absence, rather than the quantity, of a target microorganism and cannot distinguish between nonviable and viable cells. To deal with these limitations several modifications of standard PCR have recently been developed. Real-time quantitative PCR (qPCR), such as the TaqMan system, relies on the release and detection of a signal after cleavage of a fluorescent-labeled probe by the 5′–3′exonuclease activity of Taq DNA polymerase. A fluorogenic oligonucleotide probe, which specifically binds within the two PCR primer regions, is degraded in each PCR cycle, thereby monitoring the amplification of the target gene by an increasing fluorescence signal in real-time. The signal is related to the amount of amplicon present during each cycle and will increase as the amount of specific amplicon increases.66,78,79,85,86 The specificity of the probe introduces a high level of confidence concerning the detection of target microorganisms in mixed samples. Another approach with known advantages is hybridization of whole bacterial cells with fluorescent oligonucleotides (FISH) targeted to broad-range sequences (16 or 23S rDNA) or specific genes.80–82 This method is rapid and easy-to-perform and allows the in situ observation of the abundance, spatial distribution and morphology of bacterial cells.82 FISH assays can significantly shorten time to retrieve results and to initiate control measures in the food industry. Microarray technology is also a powerful tool that can be used for simultaneous detection of large sets of different genes of DNA (or RNA) targets on a glass slide.66 By

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processing multiple samples and analyzing up to tens of thousands of genes in a single assay, this method is a promising alternative for monitoring environmental and food bacteria, but it requires expensive microarrayer and laser-scanner apparatus. When applying microarrays for microorganism detection, PCR amplification of target genes it often involved in order to enhance detection sensitivity.66,87–89 Microarray methodology has already been applied for the detection of enterococci in environments such as wastewater66 and artificially contaminated milk samples.84 A summary of available probes that can be used in qPCR, RT-qPCR, hybridization assays, FISH, and microarrays for the detection, and/or quantification of enterococci is presented in Table 12.1. As stated before, a variety of molecular methods have been developed for enterococcal detection and identification at genus or species level. The detected DNA targets include genes encoding for rRNA66,74,78,79,90,91 the alpha subunit of RNA polymerase and the phenylalanyl-tRNA synthase,92 the elongation factor EF-Tu,73 the D-alanine:D-alanine ligase,70 a penicillin binding protein,67 the muramidase,77 the alpha subunit of the bacterial ATP synthase,93 the manganese dependent superoxide dismutase,72,94 and the heat shock proteins (groESL),66,76,95,96 as well as genus or species-specific antimicrobial determinants.70,71,79 A short description of each method is presented below and additional information about broad-range PCR primers (designed for eubacteria and lactic acid bacteria and to be used in positive control reactions) and enterococcal PCR primers (designed for selective amplification at genus and species level) can be found in Table 12.2. Ribosomal RNA. rRNA is a universal component of bacterial ribosomes and 5S, 16S, and 23S rRNAs are present at the small (30S) and large (50S) subunits that comprise the complete active ribosome (70S). Since rRNAs are found in high copy numbers (103–104 molecules) per actively growing cell and their coding genes are usually present in more than two copies so, targeting the rRNAs or their coding genes, has the potential to increase the detection sensitivity compared to assays based on the detection of single-copy genes.97 Small subunit ribosomal RNA (16S rRNA) gene sequences are widely used as target for the development of PCR primers and/or specific hybridization probes due to their phylogenetic informational content and wide availability in databases. By comparison, although the large subunit rRNA (23S rRNA) gene contains more sequence variations than the 16S rRNA gene, and thus may provide more useful targets for microorganism identification at species or infra-species level, it has been less frequently used mainly due to the limited availability of sequence information. Since both 16S rRNA and 23S rRNA are part of the ribosome and due to evolutionary constraints there is a minimal variation within these regions. So, examining the 16S–23S rDNA intergenic noncoding region (ITS) may be necessary to distinguish very similar microorganisms. Due to the universality of rRNA genes the selection of specific primers or probes has to be criteriously performed. The use of such gene targets will be more difficult when analyzing

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Molecular Detection of Foodborne Pathogens

Table 12.1 Probes for Detection and/or Identification of Enterococci Organism Eubacteria   Gram positive bacteria Enterococcus spp.                 vanA enterococci vanC enterococci E. asini     E. avium       E. casseliflavus E. cecorum         E. columbae       E. dispar           E. durans    

Target 16S rDNA 16S rDNA 16S rDNA 16S rDNA 16S rDNAa 16S rDNA 16S rDNA 16S rDNA 23S rDNA 23S rDNA 23S rRNAb 23S rDNA vanA gene 16S rDNA 16S rDNA 16S rDNA 23S rRNA 16S rDNA ITS region 23S rRNA 23S rRNA 23S rRNA 16S rDNA 16S rDNA ITS region ITS region 23S rRNA 16S rDNA ITS region ITS region 23S rRNA 16S rDNA 16S rDNA ITS region ITS region ITS region 23S rRNA 16S rDNA ITS region ITS region

Sequence (5’–3’) EUB338-GCTGCCTCCCGTAGGAGT GTACAAGGCCCGGGAACGTATTCACC GACATAAGGGGCATGATGATTTGACGT Encl 31-CCCCTTCTGATGGGCAGG Encl 45-GGGATAACACTTGCAAAC Encl 259-GAAGTCGCGAGGCTAAGC ENC221-CACCGCGGGTCCATCCATCA ENF191-GAAAGCGCCTTTCACTCTTATGC TGGTTCTCTCCGAAATAGCTTTAGGGCTA GPL813TQb-TGGTTCTCTCCGAAATAGCTTTAGGGCTA Enc01aV-AGGTTAAGTGAATAAGGG ENC176-CAGTTCTCTGCGTCTACCTC vana3-CAACTAACGCGGCACTGTTTCCCAAT EGAC183-CAACTTTCTTCCATGCGGAAAAT CCGCATAACAATCAAAGTCGC Edil 31-CCCCCGCTTGAGGGCAGG Eas09-CGTAACATCCTATCAAAG GAAACCGCATGGTTTCGGTTTGA TACAGAAACAATTTTAAACAA(T) Eav58i-AAATGCTTACATCTCTAA Eav58-TTAGAGATGTAAGCATTT Eca58-AGCTTGTCCGTACAGGTA CTTGCTACACCGGAAGAAAATGAG Ece92-CCACTCATTTTCTTCCGG Ecec1-TGCTTTATTATTTAAAAGATCTGA(T) Ecec2-ATATTGAAAAGTAATACAAAAAAAC(T) Ece09-CACTTAAAGGTAACATCC COL-CTTGCTACACCGAAAGAAAGTGAG Ecol1-GAAGAGAAACATCAATTAATAAACCG(T) Ecol2-ACAGGWTTTGCTTTTGTTCA(T) Eco09i-GGATATTACCCTTAAGTG DIS-TAACCTACCCTCAAGCGGGGGATA Edil 137-ATGTTATCCCCCGCTTGA Edi1-CTGGATACTTGAAGAAAAGAATCAAA(T) Edi2-TCTTGAGTAAGCTTCTATC(T) Edi3-GAAGAACTGCTCAAAACCAGACC(T) Efs38-ATTCTTCACTTCCAAATT DUR-GTCGTACGCTTCTTTTTCCACC Edu1-TGACATTAGAGGATACTCTCAAGAGAAT(T) Edu2-TAATTGAAGAGTCAGGTTATGACTTTGACA(T)

Method

Reference

FISH Microarrays Microarrays Microarrays Microarrays Microarrays FISH FISH TaqMan PCR (qPCR) TaqMan PCR (qPCR) Microarrays FISH TaqMan PCR (qPCR) FISH Hybridization Microarrays Microarrays Hybridization Hybridization Microarrays Microarrays Microarrays Hybridization Microarrays Hybridization Hybridization Microarrays Hybridization Hybridization Hybridization Microarrays Hybridization Microarrays Hybridization Hybridization Hybridization Microarrays Hybridization Hybridization Hybridization

150 151 151 86 86 86 82 82 78 78 86 82 79 82 152 86 86 152 91 86 84 84 152 86 91 91 86 152 91 91 86 152 86 91 91 91 86 152 91 91

 

23S rRNA

Edr58-CTTACTCGTGTAGACAGA

Microarrays

84

 

23S rRNA

Edr58i-TCTGTCTACACGGATAAG

Microarrays

84

E. faecalis

16S rDNA

EFA1-TGATTTGAAAGGCGCTTTCGGGTGTCGCTGATGGATGGAC

Microarrays

74

 

16S rDNA

EFA2-GAAGAACAAGGATGAGAGTAACTGTTCATCCCTTGACGG

Microarrays

74

 

16S rDNA

EFA3-GAAGTACAACGAGTTGCGAAGTCGCGAGGCTAAGCTAAT

Microarrays

 

16S rDNA

CAATTGGAAAGAGGAGTGGCGGACG

TaqMan PCR (qPCR)

74 153

 

16S rDNA

TGCCGCATGGCATAAGAGTGAAAGGCGCTTTCGGGTGTC

Microarrays

90

 

16S rDNA

CAAGGACGTTAGTAACTGAACGTCCCCTGACGGTATCTAA

Microarrays

90

 

16S rDNA

GAAGTACAACGAGTCGTTAGACCGCGAGGTCATGCAAATC

Microarrays

90

 

16S rDNA

CCTCGCGGTCTAGCAGCTCGTTGTGCTT

Microarrays

151

 

16S rDNA

Enfl84-UGCACUCAAUUGGAAAGAGG

FISH

81

 

16S rDNA

efs129-CCCTCTGATGGGTAGGTT

Microarrays

86 (Continued)

161

Enterococcus

Table 12.1  (Continued) Organism

Target

Sequence (5’–3’)

Method

 

ITS region

Efs1-TATTTATTGATTAACCTTCT(T)

Hybridization

 

ITS region

Efs2-AAGAAGTGATCAAGACCCA(T)

Hybridization

  23S rRNA Efs18i-CGAAATGCTAACAACACC   23S rRNA Efa54-CAAAAACAACTGGTACAG E. faecium 16S rDNA TGATTTGAAAGGCGCTTTCGGGTGTCGCTGATGGATGGAC   16S rDNA GAAGAACAAGGATGAGAGTAACTGTTCATCCCTTGACGG   16S rDNA GAAGTACAACGAGTTGCGAAGTCGCGAGGCTAAGCTAAT   ITS region Efm1-TTTTATGAGACGATCGAT(T)   ITS region Efm2-TCTTGATCTAACTTCTAT(T)   23S rRNA Efm09i-GGATGTTACGATTGTGTG   23S rRNA Efm09-CACACAATCGTAACATCC   23S rRNA Efi58-TGACTCCTCTCCAGACTT   23S rRNA ATCATACGATCAGCCGCAGTGAATA   23S rRNA ENU140-TTCACACAATCGTAACATCCTA   23S rRNA ENU1470-GACTCCTTCAGACTTACTGCTTGG E. flavescens 23S rRNA Efl58i-TTCTACCTATACGGACAA E. gallinarum ITS region Egal-GAGTGGACAAGTTAAAGA(T)   23S rRNA Ega09-CACAACTGTGTAACATCC   23S rRNA EGA141-ATTCACAACTGTGTAACATCCTAT a not E. dispar/E. asini/E. solitarius/E. columbae/E. cecorum/E. faecalis b not E. solitarius

products with complex mixtures of bacteria making specificity analysis fundamental for validation of the method. Nowadays, several databases compiling full and partial rDNA sequences are available online and updated regularly allowing fast and easy comparison of data from unknown organisms. Among these databases are the Ribosomal Database Project II and ProbeBase. Ribosomal Database Project II Release 9.58 (Release 9, update 58) consists of 481,650 aligned and annotated 16S rRNA sequences, along with seven online analysis tools. Update 58 was released on February 6, 2008 (http://rdp.cme.msu.edu/). ProbeBase (http://www.probebase.net) is a comprehensive database containing 1317 rRNA-targeted oligonucleotide probe sequences and eight microarrays from 288 references (status February 2008), with supporting bibliographic and biological annotation, which is frequently used by researchers worldwide. Amplification of the rRNA genes (e.g., PCR or RT-PCR) and the use of probes directed to these regions (e.g., FISH or microarrays) have been widely applied for the detection and/ or identification of Enterococcus as a genus or for particular species. Since E. faecalis is the most commonly encountered species, the majority of the primers and probes are specific for its identification. RNA polymerase and phenylalanyl-tRNA synthase. In 2005 Naser92 and coworkers reported that all enterococcal species analyzed were clearly differentiated on the basis of their RNA polymerase (rpoA) and phenylalanyl-tRNA synthase (pheS) gene sequences. Strains of the same enterococcal species had at least 99% rpoA and 97% pheS sequence similarity, whereas different enterococcal species had a

Microarrays Microarrays Microarrays Microarrays Microarrays Hybridization Hybridization Microarrays Microarrays Microarrays RT-qPCR FISH FISH Microarrays Hybridization Microarrays FISH

Reference 91 91 modified from 154 86 90 90 90 91 91 modified from 154 86 86 66 82 82 86 91 86 82

maximum of 97% rpoA and 86% pheS sequence similarity, pointing to the usefulness of this gene as a target for specieslevel molecular identification. However, no species-specific primers have yet been developed or applied to this genus. Elongation factor EF-Tu. The tuf gene, encoding elongation factor EF-Tu, is involved in peptide chain formation and is an essential constituent of the bacterial genome73,98 making this region a good target for diagnostic purposes. Ke and coworkers73 reported in 1999 a PCR-based assay that targets the tuf gene and detects most enterococcal species with excellent sensitivity and acceptable specificity. D-alanine:D-alanine ligase. A PCR assay developed by Dutka-Malen and coworkers70 enabling the identification at the species level (species E. faecium and E. faecalis) and targeting the D-alanine:D-alanine ligase (ddl) gene was applied to characterize clinically relevant enterococci. Primers for species-specific PCR of E. durans and E. hirae targeting the ddl gene were also developed and validated by Knijff and coworkers.99 Penicillin-binding proteins. Penicillin-binding proteins (PBPs) are the enzymes involved in terminal stages of peptidoglycan synthesis and PBP1 and PBP5 are the prevalent PBPs. Thus, these proteins are potential targets for bacterial cell detection. The gene coding for PBP5 (pbp5) has previously been used for the development of an E. faecalis species-specific probe100 and was subsequently used in competitive PCR for quantification of nonculturable E. faecalis cells.67 Muramidase. In 2006 Arias and coworkers77 searched the GenBank for sequences to be applied for the identification of E. hirae. During this process an E. durans muramidase gene

Eubacteria

Elongation factor EF-Tu (tuf)

U2 Rev-AYRTTITCICCIGGCATIACCAT

U1 For-AAYATGATIACIGGIGCIGCICARATGGA

803

589

590 For-GGNGACGGNACNACNACNGCAACNGT

590 Rev-TCNCCRAANCCNGGYGCNTTNACNGC

600

~600

900

~1500 ~1000 92

996

16S rDNA+23S rDNA+ITS

Sm785 For+422Rev ENFE+L189

 

73

 

96

 

95

 

91

  156 160 161   66  

 

159

 

76

 

69

170 1522

74,90

1554

156 157 158

~1500 676/578 466

Reference

155 Sm785 For+422Rev with Ef according to Aymerich et al.171

Observations

320

Product Size (bp)

Chaperonin 60 H279A For-GAIIIIGCIGGIGA(TC)GGIACIACIAC H280A Rev-(TC)(TG)I(TC)(TG)ITCICC(AG)AAICCIGGIGC(TC)TT

rrn  13B For-GTGAATACGTTCCCGGGCCT 6 Rev-GGGTTYCCCCRTTCRGAAAT

23S rDNA 422 Rev-GGAGTATTTAGCCTT L189 Rev-GGTACTTABATGTTTCAGTTC ECST784 For-AGAAATTCCAAACGAACTTG ENC854 Rev-CAGTGCTCTACCTCCATCATT For-AGGAKGTTGGCTTAGAAGCAG Rev-CGCTACCTTAGGACCGTTATAGTTAC

16S rDNA For-GGATTAGATACCCTGGTAGTCC Rev-TCGTTGCGGGACTTAACCCAAC Sm785 For-GGATTAGATACCCTGGTAGTC For-GCTGGATCACCTCCTTTC Uni331 For-TCCTACGGGAGGCAGCAGT Uni797 Rev-GGACTACCAGGGTATCTAATCCTGTT For-GAGAGTTTGATYCTGGCTCAG Rev-AAGGAGGTGATCCARCCGCA 1020 For-TTAAACTCAAAGGAATTGACGG 1190 Rev-CTCACGRCACGAGCTGACGAC Ef16 For-AGAGTTTGATCCTGGCTCA Ef16 Rev-GGTTACCTTGTTACGACTTC U1 For-CCAGCAGCCGCGGTAATACG U2 Rev-ATCGG(C/T)TACCTTGTTACGACTTC

Primers (5’–3’)

Table 12.2 Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers

162 Molecular Detection of Foodborne Pathogens

112

480

Manganese-dependent superoxide dismutase (sodA) d1 For-CCITAYICITAYGAYGCIYTIGARCC d2 Rev-ARRTARTAIGCRTGYTCCCAIACRTC

1609

320+420

Elongation factor EF-Tu (tuf) Ent1 For-TACTGACAAACCATTCATGATG Ent2 Rev-AACTTCGTCACCAACGCGAAC

Chaperonin 60 GroES EntGroES For-TTAAAACCATTAGGCGATCG EntGroES Rev-CCCATNCCCATNGANGGRTCCAT

ITS region For-CAAGGCATCCACCGT Rev-GAAGTCGTAACAAGG

92

115

16S rDNA Ec-ssu1 For-GGATAACACTTGGAAACAGG Ec-ssu1 Rev-TCCTTGTTCTTCTCTAACAA

23S rDNA ECST784 For -AGAAATTCCAAACGAACTTG ENC854 Rev-CAGTGCTCTACCTCCATCATT

733

815

16S rDNA E1 For-TCAACCGGGGAGGGT E2 Rev-ATTACTAGCGATTCCGG

Genus Enterococcus

RNA polymerase a subunit (rpoA) rpoA-21 For-ATGATYGARTTTGAAAAACC rpoA-23 Rev-ACHGTRTTRATDCCDGCRCG

431

  72  

  73  

  96  

  162, 163, 164  

  78  

  97  

  149  

 

  92  

  92   92  

Phenylalanyl-tRNA synthase (pheS) pheS-21 For-CAYCCNGCHCGYGAYATGC pheS-22 Rev-CCWARVCCRAARGCAAARCC pheS-21 For-CAYCCNGCHCGYGAYATGC pheS-23 Rev-GGRTGRACCATVCCNGCHCC 494

  93  

1102

  atpA-27 Rev-CCRCGRTTHARYTTHGCYTG

  atpA-20 For-TAYRTYGGKGAYGGDATYGC

α subunit of ATP synthase (atpA)

Lactic acid bacteria

(Continued)

Enterococcus 163

 

360

444

430

941

Manganese-dependent superoxide dismutase (sodA) FL1 For-ACTTATGTGACTAACTTAACC FL2 Rev-TAATGGTGAATCTTGGTTTGG

Penicillin binding protein (pbp5) For-CATGCGCAATTAATCGG Rev-CATAGCCTGTCGCAAAAC

vanE EE1 For-TGTGGTATCGGAGCTGCAG EE2 Rev-ATAGTTTAGCTGGTAAC

vanG EG1 For-CGGCATCCGCTGTTTTTGA EG2 Rev-GAACGATAGACCAATGCCTT

  105   475

105  

105  

 

67

 

94

 

76

70

 

76

 

66

 

96

941

650

64

185

  168

~1000

 

167

 

165 166 153

 

Reference

69 ENFE + L 189

Observations

138

112

143

360

803

E. faecalis

Product Size (bp)

Iron sulfur binding protein Efis For-ATGCCGACATTGAAAGAAAAAATT Efis Rev-TCAATCTTTGGTTCCATCTCT

D-alanine/D-alanine ligase (ddl) For-ATCAAGTACAGTTAGTCTTTATTAG Rev-ACGATTCAAAGCTAACTGAATCAGT DD13 For-CACCTGAAGAAACAGGC DD13 Rev-ATGGCTACTTCAATTTCACG

Chaperonin 60 GroESL EfGroES For-GGAATTGTTCTTGCATCCGT EfGroES Rev-ACAATTAAGTATTCTACGCC For-TGTGGCAACAGGGATCAAGA Rev-TTCAGCGATTTGACGGATTG Efes For-GTGTTAAAACCATTAGGCGAT Efes Rev-AAGCCTTCACGAACAATGG

16S rDNA Efs130 For-AACCTACCCATCAGAGGG Efs490 Rev-GACGTTCAGTTACTAACG For-CGCTTCTTTCCTCCCGAGT Rev-GCCATGCGGCATAAACTG For-TACTGACAAACCATTCATGATG Rev-AACTTCGTCACCAACGCGAAC 72 For-CCGAGTGCTTGCACTCAATTGG 210 Rev-CTCTTATGCCATGCGGCATAAAC ENFE For-GTCGCTAGACCGCGAGGTCATGA

Primers (5’–3’)

Table 12.2  (Continued)

164 Molecular Detection of Foodborne Pathogens

Muramidase (mur) mur-2ed For-AACAGCTTACTTGACTGGACGC mur-2ed Rev-GTATTGGCGCTACTACCCGTATC

D-alanine/ D-alanine ligase (ddl) DuHi For-TTATGTCCCWGTWTTGAAAAATCAA Du Rev-TGAATCATATTGGTATGCAGTCCG Hi Rev-TTTTGTTAGACCTCTTCCGGA

E. hirae and E. durans

177

377

186

658

Unknown region For-TTGAGGCAGACCAGATTGACG Rev-TATGACAGCGACTCCGATTCC

1091

550

676/578

215

E. faecium

500

297

647

298

Manganese-dependent superoxide dismutase (sodA) FM1 For-GAAAAAACAATAGAAGAATTAT FM2 Rev-TGCTTTTTTGAATTCTTCTTTA

D-alanine/D-alanine ligase (ddl) For-GCAAGGCTTCTTAGAGA Rev-CATCGTGTAAGCTAACTTC FAC1 For-GAGTAAATCACTGAACGA FAC2 Rev-CGCTGATGGTATCGATTCAT

23S rDNA Ef Rev-CACACAATCGTAACATCCTA

vanD ED1 For-TGTGGGATGCGATATTCAA ED2 Rev-TGCAGCCAAGTATCCGGTAA

with 16S For according to 171

E. durans

E. hirae and E. durans E. durans E. hirae

 

 

(Continued)

 

77

  99    

 

 

172

   

94

 

 

 

105

70

85

105  

  169   71   105   170  

433

vanB For-GTGACAAACCGGAGGCGAGGA Rev-CCGCCATCCTCCTGCAAAAAA For-CATCGCCGTCCCCGAATTTCAAA Rev-GATGCGGAAGATACCGTGGCT EB3 For-ACGGAATGGGAAGCCGA EB4 Rev-TGCACCCGATTTCGTTC For-CATCGCCGTCCCCGAATTTCAAA Rev-GATGCGGAAGATACCGTGGCT

  79  

64

 

vanA vana3 For-CTGTGAGGTCGGTTGTGCG vana3 Rev-TTTGGTCCACCTCGCCA

E. faecalis and E. faecium

Enterococcus 165

Manganese-dependent superoxide dismutase (sodA) AS1 For-GCATCATGACAAGCATCACGC AS2 Rev-GGCTTTTTGCCTTCAGATAAA AV1 For-GCTGCGATTGAAAAATATCCG AV2 Rev-AAGCCAATGATCGGTGTTTTT CA1 For-TCCTGAATTAGGTGAAAAAAC CA2 Rev-GCTAGTTTACCGTCTTTAACG CE1 For-AAACATCATAAAACCTATTTA CE2 Rev-AATGGTGAATCTTGGTTCGCA CO1 For-GAATTTGGTACCAAGACAGTT CO2 Rev-GCTAATTTACCGTTATCGACT DI1 For-GAACTAGCAGAAAAAAGTGTG DI2 Rev-GATAATTTACCGTTATTTACC DU1 For-CCTACTGATATTAAGACAGCG DU2 Rev-TAATCCTAAGATAGGTGTTTG FV1 For-GAATTAGGTGAAAAAAAAGTT FV2 Rev-GCTAGTTTACCGTCTTTAACG GA1 For-TTACTTGCTGATTTTGATTCG GA2 Rev-TGAATTCTTCTTTGAAATCAG GI1 For-CTGGCTGGGCTTGGCTAGTGA GI2 Rev-ATAATCGGTGTTTTACCGTCT HI1 For-CTTTCTGATATGGATGCTGTC HI2 Rev-TAAATTCTTCCTTAAATGTTG MA1 For-GTAACGAACTTGAATGAAGTG MA2 Rev-TTGATCGCACCTGTTGGTTTT MU1 For-CAGACATGGATGCTATTCCATCT MU2 Rev-GCCATGATTTTCCAGAAGAAT PA1 For-TGGCACCAAATGCTGGCGGAA PA2 Rev-TGGTGTAGAAGTAATTTCAAG

16S rDNA Ita For-TACCGCATAATACTTTTTCTCT Ita Rev-GTCAAGGGATGAACATTCTCT

Other enterococcal species

mur-2 For-CGTCAGTACCCTTCTTTTGCAGAGTC mur-2 Rev-GCATTATTACCAGTGTTAGTGGTTG copY For-CGAGTTATCTGGACTTTAGGTCAAGC copY Rev-CATTCAATCGTTTCGACTGGCT murG For-GGCATATTTATCCAGCACTAG murG Rev-CTCTGGATCAAGTCCATAAGTGG

Primers (5’–3’)

Table 12.2  (Continued)

E. hirae

521

E. asini E. avium E. casseliflavus E. cecorum E. columbae E. dispar E. durans E. flavescens E. gallinarum E. gilvus E. hirae E. malodoratus E. mundtii E. pallens

365 368 288 371 284 284 295 284 173 98 187 134 98 160

E. italicus

E. hirae

344

323

E. hirae

521

Product Size (bp)

Observations

 

94

 

94

 

94

 

94

 

94

 

94

 

94

 

94

 

94

 

94

 

94

 

94

 

94

 

94

 

 

173

 

 

 

77

 

77

 

77

Reference

166 Molecular Detection of Foodborne Pathogens

PO1 For-TGGTTTCTGATATGGATGCGA

Rev-CGAGCAAGACCTTTAAG

For-CTCCTACGATTCTCTTG

Rev-CGCAGGGACGGTGATTTT

For-CGGGGAAGATGGCAGTAT

vanC2/C3

Rev-CGAGCAAGACCTTTAAG

For-CTCCTACGATTCTCTTG

vanC2

EC8 Rev-TAGCGGGAGTGMCYMGTAA

EC5 For-ATGGATTGGTAYTKGTAT

vanC1/C2

Rev-CTTCCGCCATCATAGCT

For-GGTATCAAGGAAACCTC

Rev-CGGCTTGATAAAGATCGGG

For-GACCCGCTGAAATATGAAG

Rev-CTTCCGCCATCATAGCT

For-GGTATCAAGGAAACCTC

vanC1

Rev-CAGCGGCCATCATACGG

For-GCTATTCAG CTGTACTC

Rev-GGAGTAGCTATCCCAGCATT

For-TCTGCAATAGAGATAGCCGC

Rev-CCCCTTTAACGCTAATACGATCAA

For-CATGAATAGAATAAAAGTTGCAATA

vanA

SU2 Rev-CCAAATGTATCTTCGATCGCT

SU1 For-TCAGTGGAAGACTTAATCGCA

SO2 Rev-AATGGAGAATCTTGGTTTGGCGTC

SO1 For-AAACACCATAACACTTATGTGACG

SA2 Rev-GTAGAAGTCACTTCTAATAAC

SA1 For-AAACACCATAACACTTATGTG

RF2 Rev-AATGGGCTATCTTGATTCGCG

RF1 For-GTCACGAACTTGAATGAAGTT

PV2 Rev-CCGAAAGCTTCGTCAATGGCG

PV1 For-TCTGTTGAGGATTTAGTTGCA

PO2 Rev-GTAATCGCTAATTTCTCTCCA

439

484

430

815/827

822

438

822

783

377

1030

173

371

371

287

173

280

E. gallinarum and E. casseliflavus/E. flavescens

E. gallinarum and E. casseliflavus/E. flavescens

E. gallinarum and E. casseliflavus/E. flavescens

E. gallinarum and E. casseliflavus/E. flavescens

E. gallinarum and E. casseliflavus/E. flavescens

E. gallinarum and E. casseliflavus/E. flavescens

E. gallinarum and E. casseliflavus/E. flavescens

Several enterococcal species

Several enterococcal species

Several enterococcal species

E. sulfureus

E. solitarius

E. saccharolyticus

E. raffinosus

E. pseudoavium

E. villorum

170

103

71

105

170

71

 

70

 

 

 

170

71

169

 

94

 

 

 

94

94

94

   

94

94

Enterococcus 167

168

(mur) with 82% homology to E. hirae was detected, primers were developed for both genes and used for multiplex PCR. This method appears to provide a rapid and accurate identification of these two very similar enterococcal species. ATP synthase. atpA codes for the alpha subunit of the bacterial ATP synthase, which functions in ATP synthesis coupled to proton transport.101 The aim of the study developed by Naser and coworkers93 was to analyze the usefulness of atpA gene sequences for the reliable identification of Enterococcus species. All species were differentiated with a maximum of 92% similarity. The intraspecies atpA sequence similarities for all species varied from 98.6 to 100%, except for E. faecium strains that showed a lower atpA sequence similarity of 96.3%. This study clearly showed that atpA provides an alternative tool for the phylogenetic study and identification of enterococci, but no genus- or species-specific primers have been described to demonstrate this potential. Manganese-dependent superoxide dismutase. In 2000 Poyart and coworkers72 sequenced the manganese-dependent superoxide dismutase gene (sodA) of 19 species of the genus Enterococcus. Since variations in their sequences appeared to be greater between species and lower within species, this gene was used to develop species-specific primers94 that can be applied to identify 21 validly published enterococcal species. Heat shock proteins. The groESL genes (also known as chaperonin 60 genes, cpn10/60 or hsp10/60), which encode 10-kDa (GroES) and 60-kDa (GroEL) heat shock proteins, are ubiquitous and evolutionarily highly conserved among bacteria.102 Goh and coworkers95 demonstrated that reverse checkerboard hybridization methodology, based on an approximately 600-bp region of the chaperonin 60 gene, can be an efficient method for the identification of Enterococcus species. Teng and coworkers96 determined the groESL genes full-length sequence of E. faecalis and used this information to develop primers specific for this species. Antimicrobial resistance. Along the years several authors used multiplex PCR to detect both the presence of vancomycin resistance determinants and identify enterococci to the species level.70,103–106 Glycopeptide resistance has been detected in several species of enterococci with seven glycopeptide resistance genotypes already described. Five of these genotypes (vanA, vanB, vanD, vanE, and vanG) are acquired mechanisms and the other two (vanC1 and vanC2/C3) are intrinsic properties. vanA and vanB are the most common resistance genotypes. The vanA genes encode proteins that confer high-level resistance to vancomycin and teicoplanin107 and have been found in several enterococcal species.108 The vanB genes confer resistance to various concentrations of vancomycin but not teicoplanin;109 vanB-type glycopeptide resistance has been described for E. faecalis and E. faecium. The vanD phenotype is characterized by resistance to moderate levels of vancomycin and to low levels of teicoplanin;110,111 vanD determinants are constitutively expressed in E. faecalis but inducible in E. faecium.110 The vanE gene is induced by low levels of vancomycin112 and has only been detected in one

Molecular Detection of Foodborne Pathogens

strain of E. faecalis. The vanC1 and vanC2/C3 genes are specific of the motile vancomycin resistant enterococci (VRE) species E. gallinarum and E. casseliflavus/E. flavescens, respectively.70 Recently, a new vancomycin resistance locus, vanG, has been detected in four E. faecalis clinical isolates with a moderated level of resistance to vancomycin and full susceptibility to teicoplanin.113

12.1.3 Why Detect and/or Quantify Enterococci? Enterococci occur in a wide variety of food products as reported above. They are very important to achieve the final organoleptic properties of those products and can also be used as probiotics. The problems with these bacteria arise when they are related to food spoilage, in particular in meat products, or because they can act as opportunistic pathogens. Studies on enterococci isolated from food have been carried out in recent years to evaluate the detrimental aspects of their presence in food products, specially in view of their wide known capacity for gene transfer, namely of antibiotic resistances and virulence factors. However, although knowing that the presence of virulence and antimicrobial resistance determinants turns the pathogenicity potential of these bacteria higher, the establishment of a direct correlation between the presence of a known enterococcal species and infection is still a matter of discussion. Concerning enterococci, the differences between clinical and food isolates in terms of the real capacity to cause disease are yet to be disclosed. As morbidity of healthy humans resulting from foodborne enterococcal infections is very low,114 all these aspects have to be carefully balanced. From all this reasoning, and because until now no one can decided if they constitute a risk to healthy consumers, methods to detect their presence and/or quantify them are needed and a review on what can implemented in a food laboratory perspective is presented below.

12.2  Methods 12.2.1 Sample Collection and Preparation 12.2.1.1 Sample Homogenization Preparation of food samples for the analysis of Enterococcus spp. by molecular methods has to start by releasing the microbial cells from the food matrix. Food matrices are very different in nature and the analysis of semi-solid, highly viscous, particulate or solid food products is always more difficult than the analysis of a liquid food product due to matrix effect. Moreover, the presence of microbial growth inhibitors, e.g., in spices or salted fishes, has to be considered. When it comes to the extraction of bacterial DNA, several problems can be encountered like differences in cell wall structure, adhesion properties of microorganisms and chemical and biological food characteristics.115 Food sample preparation has to take into account, when preparing the initial suspension and subsequent decimal dilutions, if resuscitation procedures are needed for improved recovery of stressed microorganisms resulting from food

Enterococcus

processing and storage. The procedure starts by weighing into a sterile bowl or plastic bag a mass m (g) or by measuring a volume V (ml) (minimum of 10 g or 10 ml, unless otherwise stated) representative of the test food. A quantity of diluent equal to 9 × m g or 9 × V ml is then added. The use of buffered peptone water is the most recommended initial dilution solution.116–119 Other diluents can and have been used, not necessarily for the detection of enterococci but similar procedures can envisaged for this genus, like 0.1 M Na2HPO4 buffer (pH 8.0) for fermented food,120 sterile Ringer solution for the homogenization of cheeses29 or trypticase soy broth (TSB) for the homogenization of bread slices, ground beef, green salads, and salad dressings.121 The mixture can be homogenized in peristaltic blenders (stomachers), rotary homogenizers (blenders), vibrational mixers (pulsifiers), or other equivalent systems. In certain cases mixing can be carried out using sterile glass beads.116,119,122 The usual operating time of a peristaltic homogenizer is 1–3 min and the rotary homogenizer shall operate for no more than 2.5 min at a speed between 15,000 and 20,000 rpm. The vibrational mixer should work for 0.5–1 min.123 International standards117–119 give specific instructions for the collection and preparation of samples from meat and meat products, fresh and processed fish, crustaceans, mollusks, and similar products. The use of buffered peptone water as first diluent has the advantage that the same homogenate can be used for the detection of other microorganisms, but selective media can be used as described in pre-enrichment if enterococci are the sole target. Two approaches for the extraction of DNA can be envisaged, working with or without enrichment. If the aim of the work is just to detect the presence of the microorganism in the food sample, a previous step of enrichment can be a solution to avoid false negatives; if the study aims to characterize the microbial composition of a food sample, both at qualitative and quantitative levels, then the step of pre-enrichment has to be skipped since it would change the initial balance of the existing species. 12.2.1.2 Cell Isolation and Counting without Pre-enrichment If the aim is to characterize individual strains of Enterococcus spp. existing in a food product, isolation can be performed directly from the initial dilution prepared above, with or without preparing further dilutions depending on the expected microbial concentration. In any case, 1 ml of the initial dilution or 1 ml or 0.2 ml of further dilutions can be pour plated or spread plated, respectively, into Enterococcus selective media. More than 100 selective media have been reported along the years for the isolation of enterococci from different types of samples. However, due to variation of media composition, no single media can be pointed as the golden standard for the isolation of enterococci. Doming and coworkers12 reviewed the media for isolation and counting of enterococci and reported that some media are more selective or have high recovery rates than others. KF (Streptococcus) agar seems

169

to be a good compromise for nondairy products whereas, for dairy products, citrate azide agar can be used with better results. Kanamycin aesculin azide (KAA) agar has also been a popular choice, as well as citrate azide tween carbonate agar (CATC), aesculin-bile-azide medium (ABA ou Enterococcose), Slanetz and Bartley medium (Enterococcus agar), thallous acetate tetrazolium glucose agar (TITG), streptococcus selective agar (ScS), and crystal violet azide agar (KA).17,124 Several chromogenic media have also been developed for isolation and selection of enterococci from faecal and clinical samples, water, and food samples, like Chromocult® enterococci agar, CHROMagar Orientation, Chromogenic UTI medium, CPS ID2 or Uriselect 4.125–127 For plant material the most widely used medium is SB agar.38,39 When the aim of the work is to isolate VRE, VRE Agar can be used.12,128 Media for the isolation or counting of enterococci from food are usually incubated at 35–37°C. However, when working with milk and dairy products media are usually incubated at 45°C to inhibit the growth of other microbiota. Colony counting of enterococci isolated from the different decimal dilution allows their estimation in food. Isolated colonies can have their DNA extracted after culturing in a suitable liquid media like Man Rogosa and Sharp (MRS), brain heart infusion (BHI) or yeast extract peptone (2YT) broth. 12.2.1.3  Pre-enrichment Since the amount of living cells will considerably exceed the amount of dead cells after the enrichment procedure most PCR methods for detection of foodborne pathogens include an initial step of culture-enrichment to increase target concentration and improve sensitivity. Prior enrichment can be performed with buffered peptone water as mentioned above, or by homogenization of the food sample directly with the selective broths for enterococci such as azide dextrose broth or Chromocult® Enterococci broth. 12.2.1.4 DNA Extraction for Conventional PCR International standards on methods to detect foodborne pathogens by PCR have already been published but no details for specific genus are given, therefore almost all the procedures published are based on research laboratories work.129–131 DNA extraction, before or after enrichment, has to be tested in order to detect possible PCR-inhibitory substances. DNA extraction from isolated colonies is usually easier to perform. If an in-house DNA extraction method is to be developed, it has to take into account the degradation of proteins present in the cell extract with proteases (e.g., proteinase K); precipitation of the resulting peptides with organic solvents (e.g., a mixture of phenol and chloroform) to leave the DNA in the aqueous phase; purification of the DNA solution and further concentration using ethanol precipitation in the presence of monovalent cations; collection of the precipitated DNA by centrifugation; washing of the DNA with ethanol and resuspension in buffer (e.g., Tris-EDTA buffer or Tris buffer.132 Some of the traditional extraction methods such as the guanidium thiocyanate method133 were recently modified.134

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PCR inhibitors (e.g., polysaccharides and humic acids) are particularly abundant in food samples and are not completely removed by classical extraction protocols. Several authors have tried to extract bacterial DNA from food using filtration steps (e.g., with a 11-µm nylon filter to separate food particles), centrifugation steps,115,135 food matrix lysis136 prior to classical methods, commercial kits and mixture of both, trying to balance between extraction efficiency, DNA purity and DNA suitability for amplification.115,120,137,138 Milk samples need special treatment85–139 due to their high fat content. A summary of commercial kits that can be used for bacterial DNA extraction from food samples, and as such extract DNA from enterococci, is presented in Table 12.3.

12.2.1.5 DNA Extraction for Real-time PCR Several methods have been used to replace quantification of enterococci by plate counting which is time-consuming. However, methods that have been proposed such as DNA hybridization, enzyme immunoassays and real-time PCR detect 103–104 cfu/ml target pathogen turning culture enrichment essential. Methods that separate bacteria from food matrix and concentrate them have been developed in order to use real-time PCR to estimate the number of bacteria directly from the food homogenate. Sterile dilution of known numbers of target copies can be used to set up a standard curve which is used to determine an unknown amount of DNA (absolute quantitation).

Table 12.3 Some Examples of Kits that can be used to Extract Enterococci DNA from Food Matrixes Company

Product Name

Extraction Method

Sample Type

Omega Bio-Tek, Inc., www. omegabiotek.com Takara Bio Inc., http://www. takara-bio.com Roche Applied Science, www. roche-applied-science.com  

EZNA bacterial DNA kit, Omega

Silica based columns

Bacteria

Dr. GenTLE® (from Yeast)

GenTLE solution I lysis cells and forms noncharged complex w. nucleic acids Silica based technology

Yeasts and bacteria

Glass fibers technology

 

The MagNA Pure LC DNA Isolation Kit III High Pure PCR Template Preparation Kit, PromoKine Genomic DNA Isolation Kit II, Maxwell® 16 Cell DNA Purification Kit,

Various food samples, raw material and processed food Bacteria, Fungi

  PromoCell GmbH, www. promocell.com Promega Corporation, www. promega.com     Peqlab Biotechnologie GmbH, www.pelab.de Macherey-Nagel GmbH, www. mn-net.com  

High pure PCR template preparation kit High Pure Foodproof Kit,

Wizard® SV Genomic Purification Promega System peqGOLD Bacterial DNA Kit

Magnetic based technology Silica adsorption Guanidine-detergent lysing solution Maxwell® 16 Instrument, MagneSil® Paramagnetic Particles Silica membrane technology HiBind® silica based spin-column

NucleoSpin® Tissue

Silica membrane technology

NucleoSpin®Food

Silica based technology

 

Yeast and bacterial, blood, cultured cells, mouse tail, thymus tissues Human, animal, plant, yeast, bacterial, and viral origin Tissue culture or bacterial cells   Blood and blood cells, mammalian tissues, cultured cells, bacteria Bacteria

 

NucleoBond® AXG

Anion-exchange chromatography

Tissue, bacteria, clinical samples, and forensic samples Food (plant origin), food (animal origin), bacteria Tissue, yeast, and bacteria

 

NucleoMag Tissue

Magnetic-based technology

Tissue, cells, and bacteria

Invitek GmbH, www.invitek.eu

RTP Bacteria DNA Mini Kit

Membrane absorption based isolation

Bacteria  

 

InviMag® Bacteria DNA Mini Kit

RTP® technology (Extraction Tube containing preformulated solid lysis reagents, carrier nucleic acid, and internal RNA/DNA extraction controls) RTP® technology magnetic beads

 

Invisorb Genomic DNA Kits

Carrier adsorption based isolation

Bacteria

GE Healthcare Life Sciences, www.gehealthcare.com

Illustra bacteria genomicPrep Mini Spin Kit

Spin columns prepacked with silicamembrane

Bacteria

®

 

®

Bacteria

(Continued)

171

Enterococcus

Table 12.3  (Continued) Company

Product Name

Extraction Method

Sample Type

Fermentas, www.fermentas. com

Genomic DNA Purification Kit

Lysis of the cells with a subsequent selective DNA precipitation with detergent

Cortex Biochem, Inc., www. cortex-biochem.com

MagaZorb® DNA Isolation Kit

Magnetic separation based technology

Chemicell GmbH, www. chemicell.com  

geneMAG-DNA/Bacteria

Magnetic silica beads technology

Whole blood, serum, cell cultures, bacteria, plant, and mammalian tissues, as well as from epithelial samples Whole blood, buffy coat, leukocytes, dried blood spots, cells, tissue, formalin-fixed paraffin-embedded tissue, saliva, urine, stool, hair, swabs, milk, bacteria, seminal fluid Bacteria

geneMAG-RNA/DNA

Magnetic silica beads technology

Blood, cells, bacteria, and viruses

 

mi-Bacterial Genomic DNA Kit (mi-BD100) MagPrep® Bacterial Genomic DNA Kit, Novagen

Silica-membrane technology

Bacteria

Silica-based resin with magnetic-based separation

Bacteria

MasterPure™ Gram Positive DNA Purification Kit

Ready-Lyse™ and the Gram Positive Cell Lysis Solution

MasterPure™ DNA Purification Kit SoilMaster™ Kit

Rapid desalting process1 to remove contaminating macromolecules Hot detergent lysis process combined with a simple chromatography step that removes organic inhibitors Homogenizer for grinding your sample, a column for straining Silica-based membrane

Merck KGaA, www. merckbiosciences.co.uk   EPICENTRE® Biotechnologies, www. epibio.com    

Cartagen Molecular, Inc., www.cartagen.com Genetic ID NA, Inc., www. genetic-id.com

BioMasher™

Invitrogen Corporation, www.invitrogen.com  

ChargeSwitch® gDNA Mini Bacteria Kit PureLink™ Genomic DNA Kit

 

Ivitrogen easy dna

ZyGEM Corporation Ltd, www.zygem.com MO BIO Laboratories, Inc., www.mobio.com

PrepGEM™ Bacteria

 

Fast ID Genomic DNA Extraction Kit

PowerMicrobial™ Midi DNA Isolation Kit/Maxi (for isolation of DNA from 50 ml, 100ml) UltraClean™ Microbial DNA Isolation Kit

Magnetic beads technology

  Gram positive bacteria

Wide variety of biological sources Bacterial, plant, or fungal templates

DNA, RNA, and Protein isolation Microorganisms, tissues, plants, and raw commodities to moderately and highly processed foods Gram positive, Gram negative

Silica spin-column

Blood, tissues, cells, bacteria, swabs, and blood spots  

EA 1 enzyme, can extract high quality DNA Lysis technology (bead solution, and bead beating)

Gram positive, Gram negative, mycoplasmas, capsuled bacteria Wide range of microorganisms; Gram-positive, Gram-negative, yeast, and fungi Variety of microorganisms including bacterial spores and fungal types Bacteria, fungi, algae, and actinomycetes

Lysis technology (bead solution, and bead beating)

 

PowerSoil™ DNA Isolation Kit

 

UltraClean™ Soil DNA Isolation Kit Bacterial Genomic DNA Isolation Kit Milk Bacterial DNA Isolation Kit

Bead beating, a silica membrane technology and a new humic substance/ brown color removal procedure Bead beating technology and an inhibitor removal solution Spin column chromatography using Norgen’s resin Spin column chromatography using resin

Chemagic Food Basic Kit

Magnetic beads technology

Gram negative and Gram positive bacteria Gram negative and Gram positive bacteria Bacteria from food matix

Chemagic DNA Bacteria Kit

Magnetic beads technology

Bacteria

Norgen Biotek Corporation, www.norgenbiotek.com   chemagen AG, www. chemagen.de  

Microorganisms in the soil

(Continued)

172

Molecular Detection of Foodborne Pathogens

Table 12.3  (Continued) Company

Product Name

Extraction Method

Sample Type

Qiagen, www1.qiagen.com

DNeasy Blood & Tissue Kit

DNeasy membrane

 

Generation Capture Column Kit

Unique

Bio-Rad Laboratories Inc., www.bio-rad.com

AquaPure Genomic DNA Isolation Kits

Novel salt precipitation reagent

Bio 101, Inc., www.bio101. com  

G-NOME® kit

Proprietary “salting out” procedure

FastDNA® Kit

 

FastDNA® SPIN Kit for Soil

Specific binding of DNA by GLASSMILK® Specific binding of DNA by GLASSMILK®

 

Ver mobio ultra cean soil kit

GENPOINT® AS, www. genpoint.no

BUGS’n BEADS™

Magnetic beads technology

 

Wizard® Magnetic DNA Purification System for Food

Magnetic beads

Qbiogene

GNOME® DNA Isolation Kit

Vivantis Technologies Sdn. Bhd.¸www.vivantis.com  

GF-1 Food DNA Extraction Kit

Mini-column spin technology

Plant, animal or mixed origins

GF-1 Bacterial DNA Extraction Kit

Mini-column spin technology

Gram negative or Gram positive bacteria

 

Trifunctional purification matrix and

Animal blood and tissues, and from cells, yeast, bacteria, or viruses Whole blood, bone marrow, buffy coat, body fluids, cultured cells, cell suspensions, and Gram negative bacteria   Cultured and fixed cells, gram-negative bacteria, plant and animal tissue, and whole blood and bone marrow cells Bacteria, yeast, and animal cells and tissues of all types Animal tissues and bacteria, plants, and yeast and fungi All bacteria, fungi, plants, and animals  

 

Bacterial DNA from a wide range of materials and complex solutions, namely Enterococcus Food samples; dairy products, FishShrimps, meat and poultry, egg, fruit and vegetable, composite foods   Bacteria, plant, yeast, blood cells and other mammalian cells and tissues of all types

No work has been done directly with enterococci in food but several species of bacteria have already been quantified in food and water using real-time PCR. Two step extraction of DNA for real-time PCR separate the target cells from the food matrix prior to isolation of DNA from the capture matrix:140 buoyant density centrifugation (BDC) has been used for the detection of food pathogens, centrifugation for Yersinia enterocolytica and flotation for Campylobacter jejuni,141,142 paramagnetic beads143 or filtration.144 BDC as a sample treatment prior to qPCR diminishes the risk of having false positives due to the detection of dead cells,145 eliminates part of the inhibitory substances from the food matrix, separates the microorganisms from the food particles and allows quantification of a target organism even amongst many others.146 Commercial kits can also be used.137,147 Internal controls have also been tested in PCR-based examination of food samples to monitor the detection method, from DNA extraction throughout amplification and detection.148 12.2.1.6 DNA Extraction from a Fermented Cereal-based Food (1) Food sample (1 g) is mixed with 9 ml of 0.9% NaCl solution and homogenized for 30 sec at maximum speed with a stomacher.

(2) Bacterial cells are recovered by two-step differential centrifugation, first at 800 × g for 1 min (to remove food particles) and then at 13,000 × g for 5 min. (3) DNA extraction starts with the addition of 500 µl of TESL (25 mM Tris, 10 mM EDTA, 20% sucrose, 20 mg/ml lysozyme and 20 U mutanolysin) to the cell pellet, followed by incubation at 37ºC for 60 min with gentle shaking. (4) 100 µl of proteinase K (1 mg/ml) are added and the suspension further incubated at 55ºC for 60 min. (5) 500 µl of GES reagent (5 mM guanidium thiocyanate, 100 mM EDTA, and 0.5% sarkosyl) are added and the lysate cooled on ice for 5 min. (6) 250 µl of cold 7.5 M ammonium acetate are added to the lysate, gentle shaked until the two phases are mixed and held on ice for 10 min. (7) The sample is then purified by one extraction with chloroform:isoamyl alcohol (24:1). (8) Nucleic acids are precipitated by isopropanol, washed with 70% ethanol and dissolved in 50 µl of TE with RNase (20 µg/ml), followed by incubation at 37ºC for 30 min.

173

Enterococcus

(9) The quality and size of the food DNA is checked by electrophoresis on 1% (wt/vol) agarose gel on 0.5% TBE buffer (Tris-Borate EDTA, pH 8.0). The DNA yield is quantified spectrophotometrically.

12.2.2  Detection Procedures For the detection of enterococci in food, several approaches can be applied depending on the food matrix, as well as the enterococcal species present or to be searched in the sample. An example of a suitable procedure, as presented here (see also Figure 12.1 which presents the flowchart of the multiplex-PCR based procedure for the detection and/or identification of enterococci at genus and species level), combines the method applied for the extraction of DNA from fermented cereal-based food according to Abriouel and coworkers120 and the detection/identification of enterococci based on the genus- and species-specific multiplex PCR described by Jackson and coworkers,94 enabling the detection of 21 validly published species of Enterococcus. This procedure can also be applied to other foods, after suitable modifications in the protocol of sample treatment prior to DNA extraction to adequately recover bacterial cells (see Section 12.2.1).

12.2.2.1 Genus- and Species-specific Multiplex PCR Amplification procedure consists on the preparation of seven PCR master mixes containing 3 mM MgCl2, 0.2 mM deoxynucleoside triphosphate mix, 3.5 U of high fidelity DNA polymerase and 1.25 µl (16 µM) of each primer (for sequence see Table 12.2 and for primer combination in each mix see Figure 12.1). For primers FL1, FL2, MA1, MA2, GA1, GA2, SA1, SA2, DI1, and DI2 the use of 2.5 µl of each primer is recommended. In all amplifications a genus-specific positive control primer set (E1/E2) must be added (for sequence see Table 12.2). PCR reactions are performed in a final volume of 22.5 µl (20 µl of master mix+2.5 µl of DNA). The PCR amplification program includes an initial denaturation at 95°C for 4 min, followed by 30 amplification cycles of denaturation at 95°C for 30 sec, annealing at 55°C (Groups 1, 2, 5, and 6) or 60°C (groups 3, 4, and 7) for 1 min, and elongation at 72°C for 1 min; amplification ends with an extension step at 72°C for 7 min. After PCR amplification an aliquot of 10 µl of product is electrophoresed on a 2% 1× Tris-acetate-EDTA agarose gel containing 2 µg/ml ethidium bromide. A DNA molecular size marker (preferably with a 10 bp resolution) should be

Food sample

DNA extraction

Genus-and species-specific multiplex PCR

Target species

Primers

Group 1 E1 + E2 DU1 + DU2 FL1 + FL2 FM1 + FM2 MA1 + MA2

Group 2

Group 3

Group 4

Group 5

Group 6

Group 7

E1 + E2 AV1 + AV2 CO1 + CO2

E1 + E2 CE1 + CE2 HI1 + HI2 RF1 + RF2

E1 + E2 AS1 + AS2 GI1 + GI2 PA1 + PA2 PO1 + PO2

E1 + E2 CA1 + CA2 GA1 + GA2 SO1 + SO2

E1 + E2 DI1 + DI2 PV1 + PV2 SA1 + SA2

E1 + E2 FV1 + FV2 MU1 + MU2 SU1 + SU2

E. durans 295 bp

E. casseliflavus 288 bp

E. dispar 284 bp

E. flavescens 284 bp

E. avium 368 bp

E. cecorum 371 bp

E. asini 365 bp

E. faecalis 360 bp

E. gallinarum 173 bp

E. pseudoavium 173 bp

E. mundtii 98 bp

E. columbae 284 bp

E. hirae 187 bp

E. gilvus 98 bp

E. faecium 215 bp

E. solitarius 371 bp

E. saccharolyticus 371 bp

E. sulfureus 173 bp

E. raffinosus 287 bp

E. pallens 160 bp

E. malodoratus 134 bp

E. villorum 280 bp

Figure 12.1  Flowchart of the multiplex-PCR based procedure for detection and/or identification of enterococci at genus and species level.

174

used as standard. Following electrophoresis, the agarose gels are visualized and photographed under UV light. 12.2.2.2 Data Analysis When coupled with the genus-specific primers described by Deasy and coworkers,149 the multiplex PCR for the superoxide dismutase (sod) gene94 provides a rapid and reliable method for the detection and/or identification of the majority of enterococcal species. Using the molecular size marker included in the agarose gel, the size of the amplicons can be estimated and compared with the expected ones (Table 12.2 and Figure 12.1). If the obtained fragments show the expected molecular size for the genus and species primer set, the presence of the target species is detected in the sample (or the isolate is identified at species level if working with pure cultures isolated from food samples). When only the genus-specific amplicon is visualized, the presence of unidentified enterococci is inferred. If no amplicons are observed, with the exception of the positive control reaction tube (food sample artificially contaminated with enterococci, which demonstrates the absence of PCR inhibitors), this means that either no enterococci are present in the analyzed sample at detectable levels or the existing strains show sequence variations in sod gene that prevent primer annealing (the false negative problem can occur despite the applied methodology and has to be taken in account). Although E flavescens (multiplex group 4) and E. casseliflavus (multiplex group 2) are distinguished, these two species are now considered the same with intraspecific variation of the sodA gene sequence, thus explaining why some strains give positive amplification with both primer sets while others show positive results only for one. For other species more recently described, no primer sets were developed since no sodA gene sequences are yet available in the databases.

12.3 Conclusions and Future Perspectives To start the discussion and the conclusion on the detection and quantification of enterococci in food, and on emerging pathogens, a definition of what is a foodborne emerging pathogen is needed. According to Morse174 and Yasmine and Adams175 an emerging pathogen is an agent “causing illnesses that have recently appeared or been recognized in a population, or that are well recognized but are rapidly increasing in incidence or geographic range.” Thinking about what was presented in the first part of this chapter, the conclusion is that this definition can be applied to Enterococcus genus. Why then the precaution with which this genus is always spoken about, because it is a very important part of some food microbiota, that is responsible for the special organoleptic properties of some fermented foods, and also because if well studied and characterized enterococci can be used as probiotics. Its presence in those food products is indispensable and as such it depends on the laboratories for evaluation of the potential risks of consumption of those microorganisms. A long road

Molecular Detection of Foodborne Pathogens

has yet to be followed to avoid been simplistic and preclude, because of all virulence factors that are detected and the presence the antibiotic resistance factors that are reported, the temptation of avoiding eating the food products that have them, avoiding using them as probiotics or even, in the future, avoiding using them as starter culture for instance for cheese production. Nevertheless, efforts should be done on eliminating antibiotic-resistant enterococci from foods by reducing veterinary use of antibiotics and making a prudent use of antibiotics in humans. The lack of an established congruence between virulence determinants and demonstrated pathogenicity show that research on this subject has to continue and is continuing, especially on the aspects of the relationships between host and microbial agent.176–179 Special attention has to be given to routes of transmission of food products namely the survival of strains to the gastrointestinal traffic and their capacity to establish themselves in the human tract. Other routes of transmission have to be investigated, namely fecal contamination of water and plants by food animals and waste water, especially hospital waste water.114 All these evaluations have to take on board not only healthy people, which seem to be unaffected by enterococcal strains, but also the elderly population, infants and immunecompromised people. The microbiological risk assessment of emerging pathogens has to be performed and global data indicates that epidemiology of foodborne diseases is changing, due to changes in food industry, lifestyles, improved surveillance and recovery/detection methods.180,181 As the emergence of foodborne pathogens depends on the laboratory ability to detect, identify and recognize those agents, therefore for the case of Enterococcus a detailed combination of genus and species more specific method is proposed here based on the work of Abriouel and coworkers120 and Jackson and coworkers.94 PCR methods in general, and real-time PCR methods in particular, are increasingly used for the detection and quantification of pathogens in food samples. Two questions still remain to be answered and constitute the future research needed in this area. The first question is the case of false positive sample detection due to the simultaneous detection of dead and live cells, a nonexisting question in conventional methods using pre-enrichment or not. This question has already been approached by several authors when reporting food sample preparation, but a more detailed study for enterococci is needed, as well as a more consistent study on the effect of the different food matrixes on the capacity of detection of the methods. This question is closed related with quantification problems. Again a sound correlation between enterococcal colony forming units (cfu/ml) and concentrations obtained by real-time PCR have to be developed. In what concerns microbiology laboratories that perform food control, validation of the methods is needed as no international standard is approved for Enterococus detection and/ or quantification. Another aspect that has to be taken in consideration is the constant need to have new methods to detect/quantify, if considered necessary, all the new species that are constantly appearing in the literature. All these

Enterococcus

new routes of needed research only make the area of work with Enterococcus more challenging and on the top of food research priorities.

Acknowledgments The authors acknowledge the partial funding from different projects on Enterococcus or food quality and safety that have financed their research on the subject, namely Fundação para a Ciência e Tecnologia through Program PRAXIS XXI, project 2/2.1/BIO/1121/95 and Program SAPIENS, project POCTI/AGR/36165/99 and EU, through Specific Supported Action projects SELAMAT no CT-2004-506386 and ALCUEFOOD na 007176. T. Semedo-Lemsaddek thanks the financial support of the FCT Pos-Doctoral fellowship SFRH/ BPD/20892/2004.

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178 116. Anonymous. ISO 6887-1:1999. Microbiology of food and animal feeding stuffs – Preparation of test samples, initial suspension and decimal dilutions for microbiological examination – Part 1: General rules for the preparation of initial suspension and dilutions. 117. Anonymous. ISO 6887-2:2003. Microbiology of food and animal feeding stuffs – Preparation of test samples, initial suspension and decimal dilutions for microbiological examination – Part 2: Specific rules for the preparation of meat and meat products. 118. Anonymous. ISO 6887-3:2003. Microbiology of food and animal feeding stuffs – Preparation of test samples, initial suspension and decimal dilutions for microbiological examination – Part 3: Specific rules for the preparation of fish and fishery products. 119. Anonymous. ISO 6887-4:2003/Cor. 1:2004. Microbiology of food and animal feeding stuffs – Preparation of test samples, initial suspension and decimal dilutions for microbiological examination – Part 4: Specific rules for the preparation of products other than milk and milk products, meat and meat products, and fish and fishery products. 120. Abriouel, H., et al. Culture-independent analysis of the microbial composition of the African traditional fermented foods poto poto and degue by using three different DNA extraction methods. Int. J. Food Microbiol., 111, 228, 2006. 121. Heller, L.C., et al. Comparison of methods for DNA isolation from food samples for detection of Shiga toxin-producing Escherichia coli by real-time PCR. Appl. Environ. Microbiol., 69, 1844, 2003. 122. Anonymous. ISO 8261:2001. Milk and milk products–General guidance for the preparation of test samples, initial suspensions and decimal dilutions for microbiological examination. 123. Anonymous. ISO 7218:2007. Microbiology of food and animal feeding stuffs – General requirements and guidance for microbial examination. 124. Reuter, G. and Klein, G. Culture media for enterococci and group D-streptococci. In Handbook of Culture Media for Food Microbiology, Corry, J.E.L. et al. Eds. Elsevier Sciences B.V, Amsterdam, the Netherlands, 2003. 125. Aspevall, O. et al. Performance of four chromogenic urine culture media after one or two days of incubation compared with reference media. J. Clin. Microbiol., 40, 1500, 2002. 126. Perry, J.D. et al. Evaluation of a new chromogenic medium, Uriselect 4, for the isolation and identification of urinary tract pathogens. J. Clin. Pathol., 56, 528, 2003. 127. Miranda, J.M. et al. Evaluation of Chromocult® enterococci agar for the isolation and selective enumeration of Enterococcus spp. in broilers. Lett. Appl. Microbiol., 41, 153, 2005. 128. Ranucci, D. et al. Microbiologial characteristics’ of hamburgers and raw pork sausages, and antibiotic-resistance of isolated bacteria. Vet. Res. Commun., 28, 269, 2004. 129. Anonymous. ISO 22174:2005. Microbiology of food and animal feeding stuffs – Polymerase chain reaction (PCR) for the detection of food-borne pathogens – General requirements and definitions. 130. Anonymous. ISO 20837:2006. Microbiology of food and animal feeding stuffs – Polymerase chain reaction (PCR) for the detection of food-borne pathogens – Requirements for sample preparation for qualitative detection. 131. Anonymous. ISO 20838:2006. Microbiology of food and animal feeding stuffs – Polymerase chain reaction (PCR) for the detection of food-borne pathogens – Requirements for amplification and detection for qualitative methods.

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Enterococcus 150. Amann, R.I., Krumholz, L. and Stahl, D.A. Fluorescentoligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. J. Bacteriol., 172, 762, 1990. 151. Jin, L.Q. et al. Detection and identification of intestinal pathogenic bacteria by hybridization to oligonucleotide ­microarray., World J. Gastroenterol., 11, 7615, 2005. 152. Manero, A. and Blanch, A.R. Identification of Enterococcus spp. based on specific hybridisation with 16S rDNA probe. J. Microbiol. Methods, 50, 115, 2002. 153. Santo Domingo, J.W., Siefring S.C. and Haugland, R.A. Realtime PCR method to detect Enterococcus faecalis in water. Biotechnol. Lett., 25, 261, 2003. 154. Betzl, D., Ludwig, W. and Schleifer, K.H. Identification of lactococci and enterococci by colony hybridization with 23S rRNA-targeted oligonucleotide probes. Appl. Environ. Microbiol., 56, 2927, 1990. 155. van De Klundert, J.A.M. and Vliegenthart, J.S. PCR detection of genes coding for aminoglycoside-modifying enzymes. In Diagnostic Molecular Microbiology, Persing, D.H., et al. Eds. ASM Press, Washington, D.C., 1993. 156. McClellan, D.L. Griffen, A.L. and Leys, E.J. Age and prevalence of Porphyromonas gingivalis in children. J. Clin. Microbiol., 34, 2017, 1996. 157. Berthier, F. and Ehrlich, S.D. Rapid species identification within two groups of closely related lactobacilli using PCR primers that target the 16S/23S rRNA spacer region. FEMS Microbiol. Lett., 161, 97, 1998. 158. Nadkarni, M.A. et al. Determination of bacterial load by realtime PCR using a broad-range (universal) probe and primers set. Microbiol., 148, 257, 2002. 159. Lu, J. et al. Use of PCR with universal primers and restriction endonuclease digestions for detection and identification of common bacterial pathogens in cerebrospinal fluid. J. Clin. Microbiol., 38, 2076, 2000. 160. Kumar, P.S. et al. New bacterial species associated with chronic periodontitis. J. Dent. Res., 82, 338, 2003. 161. Volkmann, H., et al. Evaluation of inhibition and cross­reaction effects on real-time PCR applied to the total DNA of wastewater samples for the quantification of bacterial antibiotic resistance genes and taxon-specific targets. Mol. Cell Probes, 21, 125, 2007. 162. Barry, T. The 16S/23S ribosomal spacer region as a target for DNA probes to identify eubacteria. PCR Methods Appl., 1, 51, 1991. 163. Jensen, M.A., Webster, J.A. and Straus, N. Rapid identification of bacteria on the basis of polymerase chain reaction-amplified ribosomal DNA spacer polymorphisms. Appl. Environ. Microbiol., 59, 945, 1993. 164. Molander, A. et al. A protocol for polymerase chain reaction detection of Enterococcus faecalis and Enterococcus faecium from root canal. Int. Endodont. J., 35, 1, 2002. 165. Meier, H. et al. Detection of enterococci with rRNA targeted DNA probes and their use for hygienic drinking water control. Water Sci. Technol., 35, 437, 1997.

179 166. Bartosch, S. et al. Characterization of bacterial communities in feces from healthy elderly volunteers and hospitalized elderly patients by using real-time PCR and effects of antibiotic treatment on the fecal microbiota. Appl. Environ. Microbiol., 70, 3575, 2004. 167. Siqueira, J.F. and Rocas, I.N. Polymerase chain reaction-based analysis of microorganisms associated with failed endodontic treatment. Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 97, 85, 2004. 168. Gomes, B.P.F.A. et al. Enterococcus faecalis in dental rroot canals detected by culture and by polymerase chain reaction analysis. Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 102, 247, 2006. 169. Clark, N.C. et al. Characterization of glycopeptide-­resistant enterococci from U.S. hospitals. Antimicrob. Agents Chem­ other., 37, 2311, 1993. 170. Miele, A., Bandera, M. and Goldstein, B.P. Use of primers selective for vancomycin resistance genes to determine van genotype in enterococci and to study gene organization in VanA isolates. Antimicrob. Agents Chemother., 39, 1772, 1995. 171. Aymerich, T. et al. Microbial quality and direct PCR identification of lactic acid bacteria and nonpathogenic Staphylococci from artisanal low-acid sausages. Appl. Environ. Microbiol., 69, 4583, 2003. Erratum in: Appl. Environ. Microbiol., 71, 1674, 2005. 172. Cheng, S. et al. A PCR assay for identification of Entero­ coccus faecium. J. Clin. Microbiol., 35, 1248, 1997. 173. Fortina, M.G. Rapid identification of Enterococcus italicus by PCR with primers targeted to 16S rRNA gene. Lett. Appl. Microbiol., 44, 443, 2007. 174. Morse, S.S. Factors and determinants of disease emergence. Rev. Sci. Tech., 23, 443, 2004. 175. Motarjemi, Y. and Adams, M. Introduction. In Emerging Foodborne Pathogens, Motarjemi, Y. and Adams, M., Eds. Woo­dhead Publishing Limited and CRC Press LLC, England, 2006. 176. Park, S.Y. et al. Extracellular gelatinase of Enterococcus faecalis destroys a defense system in insect hemolymph and human serum. Infect. Immun., 75, 1861, 2007. 177. Brinster, S. et al. Enterococcal leucine-rich repeat containing protein involved in virulence and host inflammatory response. Infect. Immun., 75, 4463, 2007. 178. Hébert, L. et al. Enterococcus faecalis constitutes an unusual bacterial model in lysozyme resistance. Infect. Immun., 75, 5390, 2007. 179. Shimada, T. et al. Effects of lysed Enterococcal faecalis FK-23 on allergen-induced serum antibody responses and active cutaneous anaphylaxis in mice. Clin. Exp. Allergy, 34, 1784, 2004. 180. Brown, M. and McClure, P. Microbiological risk assessment for emerging pathogens. In Emerging Foodborne Pathogens, Motarjemi, Y. and Adams, M. Eds. Woodhead Publishing Limited and CRC Press LLC, England, 2006. 181. Skovgaard, N. New trends in emerging pathogens. Int. J. Food Microbiol., 120, 217, 2007.

13 Helicobacter Norihisa Noguchi

Tokyo University of Pharmacy and Life Sciences

Contents 13.1 Introduction.....................................................................................................................................................................181 13.1.1 Morphology and Classification of Helicobacter . .............................................................................................181 13.1.2 Biology, Pathogenesis, and Medical Importance.............................................................................................. 182 13.1.3 Current Technologies for Detection of Helicobacter........................................................................................ 183 13.1.3.1 Immunological Techniques.............................................................................................................. 183 13.1.3.2 Urea Breath Test (UBT)................................................................................................................... 184 13.1.3.3 Molecular Techniques...................................................................................................................... 184 13.2 Methods.......................................................................................................................................................................... 186 13.2.1 Reagents, Supplies, and Equipment.................................................................................................................. 186 13.2.2 Sample Collection and Preparation.................................................................................................................. 187 13.2.3 Detection Procedures........................................................................................................................................ 188 13.3 Conclusions and Future Perspectives............................................................................................................................. 195 References.................................................................................................................................................................................. 196

13.1  Introduction 13.1.1 Morphology and Classification of Helicobacter Although it was once considered difficult for microbes to live in the stomachs of mammals, including humans, spiral-shaped bacteria have been detected in the gastric mucosa of some mammalian species. In 1982, Warren, and Marshall were the first to isolate a spiral-shaped Gram-negative bacterium, Helicobacter pylori, from patients with chronic gastritis.1 This organism was classified as Campylobacter pylori, which was transferred to a new genus, Helicobacter, and renamed Helicobacter pylori based on taxonomies developed using 16S rRNA sequences.2 Scanning electron microscopy revealed H. pylori to be a spiral-shaped bacterium with two to three turns and four to eight polar flagella (Figure 13.1a).3,4 Helicobacter isolated from gastric mucosa has strong urease activity. Urease, an enzyme that catalyzes the hydrolysis of urea to yield ammonia, is necessary for H. pylori to maintain a microenvironment of neutral pH around the bacterium.5 H. pylori is a pathogenic bacterium that causes peptic ulcers and chronic gastritis and is linked to mucosa-associated lymphoid tissue lymphoma.6–8 This has prompted the development of several methods for diagnosing H. pylori infection (Table 13.1).9 Spiral-shaped bacteria have also been found in the stomachs of some animals. The discovery of Helicobacter and the development of culture techniques made it possible to isolate spiral-shaped Helicobacter species other than H. pylori from animals such as cats, dogs, monkeys, and cheetahs.4 H. pylori is isolated from the upper gastrointestinal tract, including the

stomach, while other Helicobacter has been isolated from not only the stomach but also the intestinal tract and feces in animals. Morphological features, such as the form of flexispira, the presence or absence of periplasmic fibers, and the number and locations of flagella, differ among Helicobacter ­species. For instance, H. mustelae, which was isolated from the stomach of a ferret, is a slightly twisted, small rod form with multiple flagella at both poles; in contrast, H. felis, which was isolated from the stomach of a cat, has multiple flagella and periplasmic fibers at the bipoles.4 To identify these morphological features, micrographs of high resolution are needed. Helicobacter isolated from the stomach has urease activity, while some species of Helicobacter isolated from the intestinal tract and liver have no urease activity. In addition to their morphological differences, Helicobacter isolates from animals have 16S rRNA and 23S rRNA sequences that are specific to each species. Therefore, the ribosomal sequences can be used for the identification of each Helicobacter species. Analyzing the similarity of the 16S rRNA gene sequences, one of the molecular methods for classifying bacterial species, can be accomplished relatively easily through DNA sequencing. However, morphological and biochemical characterizations are usually necessary to identify Helicobacter species with certainty, because the DNA sequences of ribosomal RNA differs only slightly among Helicobacter ­species. Isolation by culturing is the gold standard in bacterial detection; however, DNA sequencing of the 16S rRNA and 23S rRNA genes is useful, because such isolation by culturing is difficult for most Helicobacter species. Currently, the genus Helicobacter comprises at least 31 species that inhabit defined sites within their hosts (Table 13.2).10 181

182

Molecular Detection of Foodborne Pathogens

(a) (b)

3.8 µm 2 µm

Figure 13.1  (a) Scanning electron micrograph of Helicobacter pylori (b) and the coccid cells.

Table 13.1 Methods for Detecting Helicobacter Method

Invasiveness

Culture

Invasive Noninvasive Invasive Noninvasive Invasive

Samples Biopsy, tissue Feces Biopsy Expiration Blood

Target

Bacteria Bacteria Urease test Urease UBT Urease ELISA Helicobacter antibodies, CagA antibodies Noninvasive Urine, saliva Helicobacter antibodies Feces Helicobacter ImmunoNoninvasive Urine Helicobacter antibodies chromatograpy Feces Helicobacter PCR Inversive Biopsy, gastric juice Genes for 16S rRNA, 23S rRNA, urease, CagA, etc. Noninvasive Feces, oral cavity Genes for 16S rRNA, 23S rRNA, urease Abbreviations:   UBT, urea breath test; ELISA, enzyme-linked immunosorbent assay.

13.1.2 Biology, Pathogenesis, and Medical Importance Helicobacter is a microaerophilic bacterium, and is cultured on medium supplemented with blood and serum because of its high auxotroph. Cyclodextrin may be used as the supplement instead of blood, and brucella broth with cyclodextrin (0.1–1%) is reported to be useful for culturing H. pylori.11,12 However, whether these conditions apply to all Helicobacter species is controversial. Generally, the growth of H. pylori is better on agar than in liquid media. Helicobacter usually colonize in the digestive organs, such as the stomach, liver, and intestinal tract, and they are excreted in feces. H. pylori, which is found in gastric mucosa as a rod-shaped spiral, transforms into a coccoid form in feces (Figure 13.1b). Although isolation by culturing H. pylori from feces has been reported, the coccoid form of H. pylori is very difficult to culture. Consequently, isolating H. pylori by culturing from feces is impractical.13 Nevertheless, some Helicobacter species that colonize in the intestinal tract and liver can be cultured from feces (Table 13.2).

Detection Agent Bacteria Bacteria NH3 13CO , 14CO 2 2 IgG

Reference

IgG Helicobacter antigens IgG Helicobacter antigens DNA

4,9 42,63,85,99,100 27,101,102 28,29,103 104–106 107 108–110 24 111 23,112 40,82,113,114

DNA

42,60,71,80,87,88

As discussed above, however, it is difficult to isolate Helicobacter strains by culturing, and this process requires microbiological expertise. Rapid identification of a Helicobacter bacterium relies on detecting a protein that is specific to the bacterium in question. Gastric Helicobacter, which grow in an acidic environment, produce urease, while most enterohepatic Helicobacter do not (Table 13.2). Thus, urease is not used as a target in the detection of Helicobacter species. Because Helicobacter species other than H. bilis produce catalase, an antibody specific to H. pylori catalase may be useful for species detection and identification. Helicobacter isolated from the stomachs of animals, such as H. mustelae, H. felis, H. heilmannii, and H. acinonychis, are known to be associated with gastritis and gastric cancer, similar to H. pylori. Most of these bacteria have been isolated from lesions associated with gastritis (Table 13.2). However, pathogenetic factors, such as VacA and CagA in H. pylori, have not been found in H. felis.14 In gastric Helicobacter ­species, as well as H. pylori, the production of urease and flagella is associated with pathogenicity. H. heilmannii15 and H. felis16 have been isolated from humans, as a result of

183

Helicobacter

Table 13.2 Helicobacter Species and Hosts Helicobacter Species H. acinonychis H. anseris H. aurati H. baculiformis H. bilis H. bizzozeronii H. brantae H. canadiensis H. canis H. cetorum H. cholecystus H. cinaedi H. cynogastricus H. equorum H. felis H. fennelliae H. ganmani H. hepaticus H. marmotae H. mastomyrinus H. mesocricetorum H. muridarum H. mustelae H. nemestrinae H. pametensis H. pullorum H. pylori H. rodentium H. salomonis H. trogontum H. typhlonicus

Host Cheetah Goose Hamster Cat, human Mouse, rat, hamster, dog, cat, sheep, fox, human Dog, cat, human Goose Human, goose Dog, cat, human Dolphin, whale Hamster Human, hamster, dog, cat, fox, rat Dog Horse Cat, dog, cheetah, human Human Mouse Mouse Cat, marmot Mouse Hamster Mouse, rat Ferret Macaque Bird, pig, cat Bird Human, nonhuman primate, dog Mouse Dog, human Rat, pig, sheep Mouse

infections that may have been mediated by pets. H. hepaticus, H. canis, H. muridarum, and H. bilis have been isolated from the intestinal tracts and livers of animals. These Helicobacter species are linked with gastroenteritis, inflammatory bowel disease, and hepatitis. These enterohepatic species may be transferred from the intestinal tract to the liver by bacterial translocation. H. cinaedi,17,18 H. fennelliae,19 H. canis,20 and H. pullorum21 have been isolated from patients with diarrhea and gastroenteritis and from immune-compromised patients. In contrast to the wealth of information available for H. pylori, little is known about the infection and pathogenicity of Helicobacter species isolated from nonhuman animals. This argues for the need for further research. Another problem with identifying Helicobacter by culturing is that it usually requires sample collection by invasive procedures. Consequently, methods have been developed for detecting Helicobacter species by PCR using samples that can be collected by noninvasive processes. In this chapter, molecular methods for the detection of Helicobacter species are reviewed.

Urease

Reference

Stomach Feces Stomach Stomach Intestinal tract, liver

Site

+ + + + +

115 116 117 118 119

Stomach Feces Feces Feces Stomach Gallbladder Feces, blood Stomach Feces Stomach Feces Intestinal tract Intestinal tract, liver Intestinal tract, liver Intestinal tract, liver Feces Intestinal tract Stomach, feces Stomach Feces Intestinal tract, liver Stomach Intestinal tract Stomach Intestinal tract Feces

+ – – – + – – + – + – – + + + – + + + – – + – + + –

120 116 121 122 123 124 125 126 127 128 125 129 130 131 132 133 134 135 136 137 21 2 138 139 140 141

13.1.3 Current Technologies for Detection of Helicobacter 13.1.3.1  Immunological Techniques Two immunological methods are currently used to detect H. pylori: the detection of anti-H. pylori antibodies in blood and urine that result from H. pylori infection, and the detection of H. pylori in feces using an anti-H. pylori antibody.9,22 These methods do not require the multi-step sample preparation necessary for PCR detection. Anti-H. pylori antibodies are commercially available for use in the Latex agglutination test and enzyme-linked immunosorbent assay (ELISA). It is important to note, however, that detecting anti-H. pylori antibodies may fail to indicate H. pylori infection, because anti-H. pylori antibodies can persist in the body up to several months after the clearance of H. pylori. H. pylori in feces is detected by ELISA and immunochromatography using a monoclonal or polyclonal antibody against a surface antigen or catalase of H. pylori.23,24 Because these methods are noninvasive, highly sensitive, and specific,

184

commercial kits for detecting H. pylori using them are useful for diagnosing H. pylori infection.25 Even though feces contain a broad variety of bacterial species, commercially available anti-H. pylori antibodies are highly sensitive and specific for the detection of H. pylori. To detect Helicobacter species other than H. pylori, an antibody against the particular Helicobacter species is required. 13.1.3.2 Urea Breath Test (UBT) H. pylori produces urease, which hydrolyzes urea, that results in the production of ammonia. While some bacteria isolated from the oral cavity and stomach have urease activity, the activity of the urease from H. pylori is extremely strong.26 Consequently, urease activity, which can be detected as a pH increase due to ammonia production, has been used for the easy diagnosis of H. pylori infection.27 However, the detection of urease requires biopsies with viable H. pylori obtained by invasive procedures. Therefore, a urease breath test (UBT), which detects urease activity through a noninvasive procedure without endoscopy, was developed.28,29 In this method, 13C-labeled urea taken orally is hydrolyzed into ammonia and CO2 by the urease activity of H. pylori. As a result, 13Clabeled CO2 is absorbed into the body and released from the lungs during exhalation. Infection with H. pylori is diagnosed by measuring the amount of 13C-labeled CO2 in a breath. In the case of a positive result using this method, two possibilities should be considered: the presence of bacteria with urease activity in the oral cavity, and the presence of H. pylori with no urease activity. In fact, UBT is useful for detecting Helicobacter species other than H. pylori that also have urease acitvity.30,31 While an instrument for measuring 13C is required, UBT is generally used for the diagnosis of H. pylori infection, because this test requires no invasive procedures. 13.1.3.3  Molecular Techniques Isolation by culturing is the gold standard for the detection of Helicobacter; however, there are two major problems: the biopsies collected for culturing most Helicobacter require invasive procedures, and culturing Helicobacter is not easy. Skirrow-agar plates including polymyxin B, which are usually for the isolation of H. pylori, cannot be employed to isolate H. pullorum, because this species is susceptible to polymyxin B.32,33 Furthermore, it takes 3–10 days to detect and identify Helicobacter by culturing, although the exact time depends on the growth rate of each species. Some Helicobacter, such as H. heilmannii, has not yet been isolated by culture.34 Thus, 16S rRNA sequences data obtained by a molecular method, in addition to the bacteria’s morphological and biochemical properties, are helpful for identifying bacterial species. An immunological method for detecting H. pylori antigen in feces is commercially available, and it is a noninvasive, simple, and rapid procedure. However, the immunological reaction is specific to each Helicobacter species, so it is difficult to use in the simultaneous testing of multiple Helicobacter. In contrast, UBT can detect not only H. pylori

Molecular Detection of Foodborne Pathogens

but also other Helicobacter that have urease activity; moreover, UBT is applicable not only to humans but also to other animals, as long as it is possible to collect their breath. However, the limits of urease detection have to be defined because of the possible presence of non-Helicobacter bacteria with urease activity.35 On the other hand, molecular methods utilize the genetic information in DNA and can serve to screen simultaneously for different bacterial species. In fact, detection by PCR-based amplification of a target gene is the most general method available. PCR: Although PCR requires information about the nucleotide sequences of the target gene, PCR is able to detect the target rapidly using a small amount of DNA, i.e., a small sample of bacteria. Therefore, PCR is particularly useful for detecting bacteria that are difficult to culture, such as Helicobacter. Although the chromosomal DNA that is the temple for PCR is easily cleaved by physical forces, it is not necessary to take precautions against cleavage, because the size of the target gene is in most cases fewer than 2,000 bp. PCR can be performed on bacterial cells if the sample is confirmed to include a single bacterial strain; moreover, frozen samples as well as paraffin sections can be used. Since most samples are more stable than the biopsies containing viable H. pylori, the sensitivity of this Helicobacter detection method does not depend on how long the sample is stored or how it is transported. The following methods have been developed using PCR for the detection of Helicobacter. How specifically PCR amplifies Helicobacter target DNA depends on the primer sequences that bind to the target gene, as well as on the thermal cycling conditions. The sensitivity of PCR depends on the purity of the DNA sample and the amount of a target gene in the sample. Therefore, sequences of primers and extraction of DNA from a sample are important factors for optimal PCR. DNA extracted from bacterial cells, gastric tissue, gastric juice, dental plaques and feces can be used as a PCR template. Nevertheless, the methods for extracting DNA from gastric juice and dental plaque require further investigation because these samples frequently include PCR inhibitors, and they contain a relatively small amount of H. pylori. Target genes for detecting Helicobacter by PCR include the genes encoding urease [ureA, ureB, and glm (formally ureC)], which are detected in many Helicobacter species, and the genes encoding 16S rRNA and 23S rRNA, which are common to all bacterial species (Table 13.3). The 16S rRNA gene, which is highly conserved, is generally used for detecting bacteria including Helicobacter. Some pathogenetic genes can also serve as target genes, such as the cagA, vacA, and cdt genes. The cdt gene, which is associated with the vacuolation of the liver, differs from the vacA in H. pylori was reported to be specific to enterohepatic Helicobacter including H. hepaticus, H. bilis, H. canis, and H. pullorum.36–38 Consequently, the cdtB gene can be used effectively as the target gene to detect enterohepatic Helicobacter. The properties of PCR methods used to detect Helicobacter species are discussed as follows:

185

Helicobacter

Table 13.3 Target Genes for Detecting Helicobacter Species by PCR Helicobacter

Target Gene

H. pylori

23S rRNA 16S rRNA ureA ureB vacA

H. bilis H. bizzozeronii

H. cetorum H. equorum H. felis

H. heilmannii H. hepaticus

H. muridarum H. pullorum

cagA glmM 16S rRNA 16S rRNA ureB tRNA 16S rRNA 16S rRNA 23S rRNA 16S rRNA ureB tRNA ureB 16S rRNA 16S rRNA cdtB ureAB 16S rRNA 16S rRNA

cdtB H. rodentium H. salomonis

16S rRNA 16S rRNA ureB

H. typlonius Helicobacter spp.

tRNA 16S rRNA 16S rRNA

Accession Number

Reference

U00679 AE001439 (AE001553) AE000511(AE000644) AF507994 M60398 U05676 U29401 L11714 AAD07146 AF252626 U18766 Y09404 AF508003 –* AF292378 DQ307735 DQ307737 AY631948 AY686607 X69080 –* L25079 AF256625 L39122 AF123537 AF066862 AF066863 M80205 AF252624 L36143 L36144 AF123536 AF220065 U96297 AY368266 Y09405 AF508005 AJ130882 –* AF127912 –*

82 142 143 144,145 35 146 146 49 147 98 39 82,144 82,148 39 127 127 39 35 82,148 35 147 98 86 149 39 147 150 81 39 40,144 82,144 82,148 39 40,144

* Nucleotide sequences are available in the indicated references.

Standard PCR: Standard PCR relies on PCR for the amplification of the target gene. The amount of DNA extracted from the cells of Helicobacter bacteria and infected tissues is sufficient to allow easy amplification of the target gene. Therefore, in these cases, standard PCR can be conducted under optimal conditions. In the case of H. pylori, primers for detecting the 23S rRNA gene, which confers clarithromycin resistance, and primers for the pathogenetic genes cagA, vacA, and ureA/B have been designed. Similarly, in the case

of Helicobacter species other than H. pylori, the 16S rRNA and 23S rRNA genes and the gene encoding urease have been used as targets.39,40 When Helicobacter cells are available, colony-direct PCR can provide sufficient DNA to detect the target gene in a single amplification run. When the amplification is poor, conducting a second PCR reaction using the first PCR products as the template is also useful, as long as the possibility of a false positive is taken into consideration. The amplified DNA is analyzed by 1–3% agarose gel electrophoresis.41 Nested PCR: Nested PCR or semi-nested PCR consists of a first PCR reaction using primers with low specificity for a target gene, followed by a second PCR using primers with high specificity for the target gene. Nested PCR can be useful for analyzing DNA extracted from samples containing a small amount of Helicobacter, abundant bacteria other than Helicobacter, or substances that inhibit PCR, such as feces, gastric juice, and dental plaque. Nested PCR can detect the target gene with both high sensitivity and high specificity. Performing two rounds of low-specificity amplification (the first PCR in the nested PCR method) prior to the high­specificity amplification may also be helpful. Because of its high sensitivity, nested PCR can lead to false positives due to contamination during PCR procedures. Thus, the standard PCR is recommended; however, nested PCR is useful when samples containing a relatively small amount of Helicobacter, such as feces, are used. In these cases, a negative control should be performed in parallel to exclude the possibility of false positives. In this work, we use nested PCR in combination with UBT or a stool antigen test to diagnose infection with clarithromycin-resistant H. pylori.42 Multiplex-PCR: Multiplex-PCR is PCR using multiple sets of primers to amplify sequences of different lengths. Using multiplex-PCR, it is possible to amplify a variety of target genes in a single PCR, depending on the species and the presence of multiple genes. Multiplex-PCR is available for detecting and typing Helicobacter species, whereas the determination procedure is complicated. Real-time PCR: Real-time PCR can detect target genes readily and quantitatively. A mutation or SNP can be detected on the basis of differences in amplification rates for different amplicons. Agarose gel electrophoresis is not required to detect target genes, and it is possible to follow the amplification in real-time through the addition of fluorescent dyes that intercalate into double-stranded DNA, or through the use of TaqMan-PCR, which uses a labeling probe. Real-time PCR is useful for detecting mutations that confer clarithromycin resistance in H. pylori, as well as for detecting other Helicobacter species. Real-time PCR requires specific equipment. RT-PCR: PCR detects not only viable bacteria but also dead bacteria because it works at the level of DNA. In contract, it is difficult to detect mRNA from dead bacteria, since mRNA turnover is extremely rapid. Thus, RT-PCR using reverse transcriptase is useful for detecting viable Helicobacter species.43–46 PCR-RFLP: PCR-RFLP is a method for identifying bacteria based on the pattern of restriction fragments generated

186

by incubating PCR-amplified DNA with restriction enzymes. The usefulness of PCR-RFLP has been demonstrated for the detection of clarithromycin-resistant H. pylori based on mutations in the 23S rRNA gene.47 PCR-DGGE: PCR-DGGE uses denaturing gradient gel electrophoresis (DGGE) to analyze the PCR product.48 The gel, which contains a gradient of the DNA-denaturing reagent urea or formamide, can resolve amplicons identical in length but different in nucleotide sequence. The difference in the hydrogen-bounding strengths in A-T and G-C base pairs leads to the separation of fragments with different nucleotide sequences. In addition, amplification fragments containing GC-rich sequences (GC clamps) migrate very slowly because of the strong binding between GC clamps. PCR-DGGE is used to identify bacterial species based on sequence differences in 16S rRNA genes. LAMP: Loop-mediated isothermal amplification is a method for amplifying a target gene at 65oC using four types of primers.49,50 Unlike PCR, LAMP does not require a thermal cycler, and it is possible to amplify a target gene efficiently. The amplified DNA is then detected by agarose gel electrophoresis. A LAMP method targeting the glm gene in H. pylori has been used to detect this H. pylori species.49 Southern hybridization: Hybridization of radiolabeled DNA probes to target DNA has long been used to identify target sequences, and, more recently, a method using nonradio labeled probes has been developed.41 The same target gene as for PCR can be used as the DNA probe. Thus, the DNA probe is prepared using PCR. Southern and colony hybridizations can be applied as methods to detect Helicobacter species. Because the hybridizations require complicated procedures, PCR is usually preferred for its ease and high sensitivity. Nevertheless, fluorescence in situ hybridization (FISH) may be useful for detecting Helicobacter species in formalin-fixed samples.51–54

13.2  Methods 13.2.1 Reagents, Supplies, and Equipment The reagents, supplies, and equipment used to detect Helicobacter are listed in Table 13.4. When samples are collected and handled, the risk of infection with Helicobacter has to be considered. Thus, the samples should be handled in a safety cabinet until the extraction of DNA is complete. Certain molecular biology supplies, such as tips and tubes, should be autoclaved before use. To avoid contamination, the PCR tubes should be handled using forceps and not with bare hands. It is recommended that the preparation for PCR also be performed in the safety cabinet to avoid aerosol contamination. Sterilized, ultra-pure water should be used for PCR. Because many formulations of Taq polymerase are commercially available with different characteristics, the most suitable type should be selected for the particular detection method. For example, in cases of detection using cultured bacterial cells, such as colony-direct PCR, premixed PCR reagents are sufficient. In cases of detection using

Molecular Detection of Foodborne Pathogens

Table 13.4 Reagents, Supplies, and Equipment in the Present Works Materials

Reference

Reagent Highly purified H2O Taq polymerase PCR buffer dNTP mixture BSA LAMP kit Oligonucleotides for primers TAE buffer Agarose Molecular marker 100-bp ladder DNA extraction kit for tissues DNA extraction kit for feces DNA purification kit Supplies Pipette tips

Plastic microtube for PCR Plastic microtube Storage tube (for extraction of DNA) Pipetter

Equipment Microcentrifuge (at up to 15,000 rpm) Tabletop microcentrifuge (at up to 3,000 rpm) Agarose electrophoresis UV transilluminator Thermal cycler Grinder

42,57,74,75,79,82,85,87,151 82,87 75 49 82 41,74,79,82,88,114 82 79,82,152 87 49,74,113

2 µl 20 µl 1000 ml 250 µl 2 ml 2 ml

42

1000 µl 200 µl 100 µl 20 µl 2 µl

48,74,80 74,82 72,85,113 42,83

samples such as gastric tissue, gastric juice, and feces, the use of Taq polymerase with high amplification efficiency is ­recommended. To reduce the emergence of unwanted amplification fragments, Taq polymerase containing an antibody (“hot-start” formulation) may be useful. The activity of this type of Taq polymerase is inhibited by the binding of the antibody at room temperature, so that false-positive reactions due to the partial annealing of DNA with primers are prevented. Once a heat-shock treatment occurs in a ­DNA-denaturing procedure, the antibody is inactivated by heating, and the Taq polymerase begins to work. This helps to ensure that

Helicobacter

only DNA that specifically anneals with primers is used as the template. The hot-start type of Taq polymerase is relatively stable at high temperatures. When detecting differences in the 16S rRNA genes in order to identify bacterial species, high fidelity Taq polymerase is recommended. The reaction buffer provided with Taq polymerase is generally optimal. Oligonucleotide primers can be obtained commercially. Bovine serum albumin (BSA) at the levels recommended for restriction enzymes and PCR should be used. A low-speed microcentrifuge is useful for collecting drops in a tube. The pipetter used for PCR should be kept separate from other pipetters. Since the time necessary to maintain a constant temperature is different for each thermal cycler, the times for denaturing, annealing, and extension should be determined specifically. The equipment for electrophoresis of the PCR-DGGE samples is available commercially.

13.2.2 Sample Collection and Preparation Biopsies, feces, gastric juices, and dental plaque can be used as samples for the detection of Helicobacter species. The use of endoscopy to biopsy tissue and collect gastric juice samples is invasive for patients, but dental plaque and feces can easily be collected without endoscopy. Cultures. As with cultures of most bacteria, DNA can easily be extracted from Helicobacter cultures. Classical phenol-chloroform extraction, boiling, and commercial kits for DNA extraction are available for obtaining sufficient DNA for PCR.55 Mixed infection with clarithromycin-resistant and ­-susceptible H. pylori has been reported previously.42,56 Therefore, it is possible for a stomach to contain different strains. For the screening of different H. pylori strains without the extraction of DNA, colony-direct PCR can be used to amplify target DNA. While it is convenient to add bacterial cells directly by using a sterilized toothpick to a tube containing PCR mixture, care should be taken to not add too many cells in one tube, which may potentially lead to the inhibition of PCR amplification. Thus, it is recommended that 1 μl of cell suspension, which is prepared by transfering the bacterial cells from one colony to 100 μl of sterilized water, be added to the PCR mixture. Biopsy. Endoscopy is used to biopsy the gastric mucosa of Helicobacter-colonized stomachs. Stomach, intestinal tract, and liver tissues obtained surgically are also used. In the case of small animals in which endoscopy is difficult, surgical resection or slaughter is required to obtain samples. The biopsies and tissues so obtained are used for detection by culturing, urease test, or PCR. For culturing, biopsies are crushed using a mortar or some other tool, and the samples are spread on a plate for the selection of Helicobacter species. Because these samples contain few substances that inhibit PCR, extracting DNA from biopsies and tissues to use in PCR is simple; these sources can be subjected to classical phenolchloroform extraction, boiling, or ­commercial-kit-based extractions to obtain sufficient DNA of adequate quality for PCR. A crushing-by-beads method is also available for DNA

187

extraction from biopsies. Frozen biopsies and tissues are also suitable for extracting DNA for use in PCR. Gastric juice. DNA extracted from gastric juice obtained by endoscopy and a string device may be used as the template for PCR.57–60 Gastric juice can be frozen until use. Gastric juice is centrifuged, and DNA is extracted from the resuspended pellet by classical phenol-chloroform extraction and boiling, in a manner similar to DNA extraction from tissue biopsies.57 Commercial kits for DNA extraction from gastric juice are also available.57,58 Feces. Feces can be obtained noninvasively from infants and small animals, for which endoscopy is difficult. Thus, feces are useful as samples for the detection of Helicobacter species. In contrast to blood and tissues, however, feces include abundant bacteria, host-derived cells such as leukocytes, and substances that inhibit PCR. As a result, the extraction of DNA from feces must be done with care to ensure the purity of the isolated DNA and to minimize contamination with substances that inhibit PCR. DNA is extracted from feces mainly through chemical methods using commercial kits, or a physical method using a cell crusher.61 Though commercial kits are easy to use and require no special instruments,62–64 DNA extraction by commercial kits is readily affected by the chemical reagents and enzymes that are used for cell lysis and for the subsequent extraction and purification of DNA. This means that, when the amount of Helicobacter DNA in the feces is small, commercial extraction kits may not provide sufficient DNA for PCR.42 To reduce this problem, a method for removing substances that inhibit PCR using an antibody against H. pylori has been developed.65 As an alternative to chemical extraction, physical methods for crushing cells in feces using beads have been developed (Figure 13.2).42,66 These methods have been used to extract DNA from microbes, including spores and mycelium, from soil.67 In these studies, approximately 50 mg of feces were added to a tube containing three types of beads of different sizes [250 mg of silica powder (63–210 µm), 32.5 mg of ceramic beads (1–2 mm), and 75 mg of glass beads], 980 µl of sodium phosphate buffer, and 180 µl of 7.5 M guanidine solution containing 5% sarcosine. Then, the suspensions were homogenized for 20–30 sec in the homogenizer. Because of the amount of debris in feces, the binding of DNA to glass beads is negligible. Following the removal of residual cells by centrifugation, DNA is purified using a commercial kit designed to purify DNA for PCR. This method usually yields 1–25 µg of DNA from 50 mg of feces. The condition of the feces affects the amount of DNA obtained; diarrheal samples, for example, often do not yield sufficient DNA. Purified DNA can be stored at –20oC for at least 1 year. In fact, it has been reported that DNA suitable for PCR amplification can be obtained from 6-yearold feces stored at –80oC.42 In a study in which controlled amounts of H. pylori were artificially added to feces, the PCR-based method was found to be capable of detecting H. pylori in feces containing ≥10 cells/50 mg of feces. The “stool crushing-by-beads” method requires a specific type of homogenizer, which can be used in the extraction of not

188

Molecular Detection of Foodborne Pathogens

Step 1: Lysis of bacteria cells in stools Beads

Stools 50 mg

Crushing 20 sec

Tube 5% Sarcosine 7.5M Guanidine

Step 2 : Extraction and purification of DNA Purification

Centrifugation

DNA H2O

Supernatant Wash

(Salt precipitation)

Figure 13.2  Isolation of Helicobacter pylori DNA from feces using the crush-by-beads method.

only DNA but also RNA and proteins, and several manufacturers sell the required type of homogenizer.68 This method can also be used to extract of DNA from tissue. Plaque. Helicobacter species have been detected in human and dog oral cavities.69 In this method, dental plaque, tongue plaque, or saliva is collected with sterile cotton swabs. The collected plaque is suspended in sterilized water and washed by centrifugation and resuspension. A portion of the cell suspension is used for culturing and the remaining suspension is boiled and used as a template in PCR.70 A commercial kit for DNA extraction from tissue can also be used to extract DNA from plaque.71 In theory at least, DNA can be extracted from fixed gastric tissue sections and checked by PCR for the presence of H. pylori.72

13.2.3  Detection Procedures Primers, target genes, and conditions for the detection of H. pylori by PCR are shown in Table 13.5. The conditions for the detection of Helicobacter other than H. pylori are summarized in Table 13.6. The annealing temperature in PCR depends on the designed primers, so these data are also presented in Tables 13.5 and 13.6. Conditions such as temperature, number of cycles, and the composition of the reaction solution for PCR depend on the Taq polymerase used. For example, Taq polymerase with an antibody (“hot-start type”) requires at least 30 cycles, because this type of Taq polymerase is fully

activated by heat only after several cycles. Alternatively, hotstart PCR can be performed by adding Taq polymerase after the denaturation at 94–95˚C and annealing at 60˚C in order to prevent nonspecific reactions due to mismatched bindings of primers at low temperatures.73 In the case of samples such as feces and plaques, which include substances that inhibit PCR, the addition of BSA at a final concentration of 0.4– 0.5% increases the amplification efficiency.74,75 Helicobacter pylori. Since many studies have reported an association of H. pylori infection with peptic ulcers and gastric cancer, numerous methods for the diagnosis of H. pylori infection have been developed. Because they do not require invasive procedures or special equipment, UBT and the ELISA method for detecting H. pylori in feces are generally preferred over molecular methods for the diagnosis of infection with H. pylori. A molecular method using PCR is employed for detecting the mutation in 23S rRNA responsible for clarithromycin resistance and the presence of pathogenetic genes such as cagA and vacA.76,77 In particular, typing of the cagA gene is used in the analysis of H. pyloriassociated gastric cancer, because a specific type of CagA has been demonstrated to be associated with gastric cancer. Since H. pylori has only one copy of the glm gene, this gene can be used as a housekeeping gene to determine the quantity of the target gene.78 Methods for the detection of H. pylori have been presented in detail in the review by Megraud and Lehours.9

Semi-nested PCR

Semi-nested PCR

Semi-nested PCR

PCR

Method

ATTCCCATTAGCAGTGCT

CEF-4

TAGGAAGTGTGAGCCGATTTG

A-2R

GTCGGTTAAATACCGACCTG AGTGGAGGTGAAAATTCC TAAGAGCCAAAGCCCTTA

GGATAAGCTTTTAGGGGTGTTAGGGG GCTTACTTTCTAACACTAACGCGC ATAATGCTAAATTAGACAACTTGAG AGAAACAAAAGCAATACGATCATTC ATATTATGGAAGAAGCGAGAGC CATGAAGTGGGTATTGAAGC

GlmMF GlmMR CagAF CagAR A-2F2 A-2F3

HP4 CRF-4 RR-1

AACGATGAAGCTTCTAGCTTGTAG GTGCTTATTCSTNAGATACCGTCAT

HeliF HeliR

CCACAGCGATGTGGTCTCAG TGTGTAGCTACCCAGCGATGCTC

GTAGGTCCTGCTACTGAAGCCTTA ATGGAAATACAACAAACACAC CTGCTTGAATGCGCCAAAC GGCCCCAATGCAGTCATGGAT GCTGTTAGTGCCTAAAGAAGCAT GGAGCCCCAGGAAACATTG CATAACTAGCGCCTTGCAC GGCAATGGTGGTCCTGGAGCTAGGC GGAAATCTTTAATCTCAGTTCGG ACATTTTGGCTAAATAAACGCTG CCAACGTGCGTAAAAGGGAATTAG GTTAARAATRGTGTRAAYGG TTTAGCTTCTGATACCGC

ureB-R VA1-F VA1-R VAm-F3 VAm-R3 VA4-F VA4-R cagA5 cagA2 Lunil 1 R5280 cagA2530S cagA3000AS

HP1 HP2

TGGGATTAGCGAGTATG CCCATTTGACTCAATG AATGCAGAAATATCAC ACTTTATTGGCTGGTTT CGTCCGGCAATAGCTGCCATAGT

HPU54 HPU18 HPU55 HPU17 ureB-F

Primer Sequence (5’–3’)

GCCAATGGTAAATTAGTT CTCCTTAATTGTTTTTAC

HPU1 HPU2

Primer

Table 13.5 PCR Protocols to Detect Helicobacter Pylori Target

cagA (5’-end) cagPAI Empty site cagA

324 bp 550 bp

68 bp

783 bp 135 bp

993 bp

204 bp

23S rRNA

23S rRNA

ureA

cagA

128 bp 314 bp

glmM

294 bp

339 bp

16S rRNA

vacA m2

352 bp

370–670 bp

vacA m1b

290 bp

ureB

464 bp vacA s1/s2

ureB

115 bp

259 bp

ureB

ureA

132 bp

411 bp

Product

Reaction

94oC30s,50oC30s,72oC1m) × 20

94oC30s,50oC30s,72oC1m) × 30 2nd (CRF-4/CRR-1):

(95oC45s,65oC45s,72oC45s) × 30,72oC4m 2nd (HP4/HP2):described above 1st (CRF-4/CRR-1):

(94oC1m,55oC1m,72oC1m) × 20 1st (HP1/HP2): 95oC2m.

94oC1m,55oC1m,72oC1m) × 30 2nd (A-2F3/A-2R):

1st (A2F2/A-2R):

(94oC30s,55oC30s,72oC3m) × 40,72oC5m

(94oC30s,50oC45s,72oC45s) × 35,72oC5m 95oC5m,

94oC5m,

(94oC1m,55oC1m,72oC1m) × 35

(94oC1m,45oC1m,72oC1m) × 35, 72oC5m

95oC5m,

Year

2005 1998

2003

1998 1996

2007 1999

2007

2005

1992

(Continued)

62 152

63

60 156

154 155

113

57

153

Reference

Helicobacter 189

CCTGARACCGTTCCTACAGC CACAGCCACTTTCAATAACGA CGTCAAAATAATTCCAAGGG GTGGATGCYCATACRGCTWA RTGAGCTTGTTGATATTGAC TTGACCAACAACCACAAACCGAAG CTTCCCTTAATTGCGAGATTCC TAGATACGGCTAATGGCG ACTAGCCTGTCAGCATCG GTTACACCCGTTAGGCTCATCACCG GTCGTTTTTAGCGAG ATCAATGCGGGGCTTTACACGCAAA GCCTAAATCTGCG

VA1XR (-) HPMGF (+) HPMGR (–) MF1 (+) MR1 (+) cagAF (+) cagAR (–) lm-F3 glm-R3 lm-FIP

glm-BIP

ATGGAAATACAACAAACACA CTGCTTGAATGCGCCAAAC

VA1F (+) VA1R (–)

AGGATGCGTCAGTCGCAAGAT CCTGTGGATAACACAGGCCAGT

Hp23S 1942F Hp23S 2308R

Primer Sequence (5’–3’)

GGTCTCAGCAAAGAGTCCCT CCCACCAAGCATTGTCCT

Primer

Hp23S 1835F H23S 2327R

107/182 bp

176/203 bp

401/475 bp

259/286 bp

367 bp

493 bp

Product

Target

glmM (ureC)

cagA

vacA m region

vacA s region

23S rRNA

Reaction

63oC60m,80oC2m

(95oC1m,50oC1m,74oC1m) × 40,74oC5m

95oC2m,(94oC10s,63oC20) × 24 95oC2m,

(94oC15s,57oC15s,72oC20s) × 30 2nd (Hp23S 1942F/Hp23S 2308R):

95oC2m,(94oC30s,57oC30s,72Co30s) × 5,

1st (Hp23S1835F/Hp23S2327R):

Notes: C, temperature in °C; m, minutes; s, seconds; ( ) ×, (reaction condition) and cycle; N=A/T/C/G, B=C/G/T, D=A/G/T, R=A/G, S=G/C, W=A/T, Y=C/T. Abbreviations:   LAMP, loop-mediated isothermal amplification; DGGE, denaturing gradient gel electrophoresis.

LAMP

PCR LiPA

Nested PCR

Method

Table 13.5  (Continued) Year

2006

1998

2006

49

146

42

Reference

190 Molecular Detection of Foodborne Pathogens

ATTCCACCTACCTCTCCCA TCTCCCATACTCTAGAAAAGT GCTTAACACATGCAAGTCGAA AGGCCCGGGAACCTATTCAC TATGACGGGTATCCGGC

ATTCCACCTACCTCTCCCA GCTATGACGGGTATCC

GATTTTACCCCTACACCA ACTTCACCCCAGTCGCTG GGGCGATAAAGTGCGCTTG

CTGGTCAATGAGAGCAGG GGAATTCCAGATCTATGAAA AAGATTAGCAGAAAAG GGAATTCGTCGACCTAGAA AATGCTAAAGAGTTG ATGAAACTAACGCCTAAAGAA CTAG GGAGAGATAAAGTGAATATGC GT GCTATGACGGGTATCC

ACTTCACCCCAGTCGCTG GTTATGTGCCTCTTAGTTTG AGAGTTCTCAGCATAACCT TGCGTAGGCGGGGTTGTAAG

CAGAGTTGTAGTTTCAAATGC

TTGGGAGGCTTTGTCTTTCCA

H676r Hbr g2 p13B F

R C97 forward

C98 reverse C05 reverse Hheilmannii F

Hheilmannii R Hpylori F

C97

C05 HceF HceR CAR577f

CAR636r

V832f

PCR

PCR

PCR

PCR

PCR

Hfelis R

Hfelis F

Hpylori R

CTGTTTTCAAGCTCCCC CTATGACGGGTATCCGGC

B39 H276f

PCR

Primer Sequence (5’–3’)

GCATTTGAAACTGTTACTCTG

Primer

B38

PCR

Method

Table 13.6 PCR to Detect Helicobacter Species

456 bp

78 bp

1,022 bp

1,200 bp

1,150 bp

1,707 bp

1,219 bp 580 bp

422 bp

375 bp

1.3 kb

374 bp

417 bp

Product

16S rRNA

16S rRNA

16S rRNA

ureB

ureB

ureB

16S rRNA

16S rRNA

16S rRNA

16S rRNA

Target Gene

H. bizzozeronii, H. salomonis H. suis

H. felis

H. cetorum

Helicobacter species

H. felis

H. pylori

H. heilmannii

Helicobacter species

Helicobacter species

H. bilis Bacteria

Helicobacter species

H. hepaticus

Target Bacteria

Samples

Reactions 94oC4m, (94oC60s,61oC135s,72oC150s) × 35, 72oC7m

94oC4m, (94oC1m,58oC2m,72oC3m) × 35, 72oC8m

94oC3m,57oC2m,72oC3m, (94oC30s,57oC30s,72oC1m) × 31, 72oC5m

94oC4m,55C, (94oC1m,55oC150s,72oC3m) × 30–40

94oC30s, (94oC2s,53oC2s,72oC30s) × 45

95oC3m, (94oC30s,60oC30s,72oC45s) × 40, 72oC5m

Culture, tissues 95oC3m, (94oC30s,60oC30s,72oC45s) × 35, 72oC5m

Feces

Gastric biospy

Tissue, bile

Feces

Culture, tissues 94oC5m, (94oC2s,53oC2s,72oC30s) × 35

Culture, feces

2004

2003

1998

1998

1997

1996

1995

Year

(Continued)

82

151

35

40

157

114

88

Reference

Helicobacter 191

GTTTGATGCGGAAGTTGTCG CAYGAYTGCACCACTTATGG TGRATTTTAAARCCAATSGC TGCGTAGGCGGGGTTGTAAG

CAGAGTTGTAGTTTCAAATGC

AGAGCGTGTAGGCGGAATGAT CGAGGAGACAAGCCCCCCGA GATTAGCTCTGCCTCGCGGCT

TTGGGAGGCTTTGTCTTTCCA GTGGAGTACAAGACCCGGGAA

CCAAGGGCTATGACGGGTGTAT CC TCTGGACTGAAGCCGGGGG

ACTTCCATCAGTTGGCTCACC CTATGACGGGTATCCGGC

ATTCCACCTACCTCTCCCA GTNGCNACBTGGAAYCTNCA

RGG DACNGGRAARTGRTC CTCCTAAGCCCACCAGAAATTG

CTTATCGCAGTCTAGTACG TTCCCCATAATAGGGTAGTTTA

UnR UvF UwR CAR577f

CAR636r

R574f R832r V1261r

V832f REVERS

HELIP2

EqF

EqR H276f

H676r VAT2

DHF1 Gan-F

16-23SR Gan-R

PCR

PCR

PCR

Semi-nested PCR

PCR

PCR

CGGATTTGATGCAAGAAGGC

UmF

Primer Sequence (5’–3’)

GATTAGCTCTGCCTCGCGGCT

Primer

V1261r

Method

Table 13.6  (Continued)

714 bp

400 bp

1,074 bp

1,107 bp

16S–23S rDNA internal spacer region

cdtB

16S rRNA

23S rRNA

H. ganmani

H. pullorum

Helicobacter species

H. equorum

Helicobacter species

Candidantus H. suis

433 bp

H. bizzozeronii, H. salomonis, H. felis

H. felis

H. felis

Target Bacteria

H. bovis

16S rRNA

16S rRNA

ure

ure

Target Gene

259 bp

78 bp

563 bp

1,770 bp

Product

Tissues

Culture

Tissues

Feces

Feces, culture

Paraffinembedded gastric biopsy specimen

Samples

94oC2m, (94oC30s,48oC30s,72oC30s) × 30, 72oC7m

94oC5m, (94oC1m,42oC2m,72oC3m) × 35, 72oC8m

95oC5m, (94oC1m,53oC1m,72oC1m) × 35, 72oC5m

94oC5m, (94oC10s,68oC40s,72oC90s) × 40, 72oC5m

94oC5m, (94oC2m,65oC1m,72oC1m) × 30, 72oC10m

95oC9m, (94oC45s,52oC45s,72oC45s) × 50, 72oC10m

95oC5m, (94oC1m,52oC1m,72oC1m) × 35, 72oC7m

Reactions

2004

2006

2007

2007

2006

2005

Year

80

89

79

87

158

72

Reference

192 Molecular Detection of Foodborne Pathogens

TaqMan PCR

Multiplex PCR

Multiplex PCR

RFLPPCR(AluI)

Nested PCR

CGTGGAGGATGAAGGTTTTA

AATTCCACCTACCTCTCCC GGGCGATAAAGTGCGCTTG

CTGGTCAATGAGAGCAGG AGGTCGCGGGTTCGAATCC

ACCAACTGGGCTAAGCGACC AACCAAYAGCCCCAGCAGCC TGGTTTTAAGGTTCCAGCGC TTTGGTGCTCACTAACGCCCTC TTCAATCTGATCGCGTAAAG TTGGGAGGCTTTGTCTTTCCA

GATTAGCTCTGCCTCGCGGCT TTGTGAAATGGAGCAAATCTT

2nd HelF

HelR2 F

R T3B(TET)

HT135R Bi1F(HEX) Bi2R Fe1F(NED) Fe3r V832f(NED)

V1261r 1201f

cdtBF

p30r

p30f

p25r

p25f

p17r

262r p17f

AAAAACT TAGCCAGTTTGGCATTCC AGGGACTCTTAAATATGCTCCT AGAGT ATTCATCGTGTTTGAATGCGTC AAATGGAACAGATAAAGATTT TAAAGCAACTTCAG CTATGCAAGTTGTGCGTTAAGC AT ATGGGTAAGAAAATAGCAAAA AGATTGCAA CTATTTCATATCCATAAGCTCTT GAGAATC ATGACAAAAAAATATTCTTTCA CAAAACTATTCATTGGT TTTATTTTAGATTCCATTTAACT GCTAAATCATCAATAGT CCGCAAATTGCAGCAATACTT

GATTTTACCCCTACACCA

C98

1375r 163f

GCTATGACGGGTATCC

1st C97

16S rRNA

447 bp

81 bp

807 bp

705 bp

435 bp

122 bp

cdtB

16S rRNA

ureB

434 bp

191 bp

ureB

tRNA

ureB

16S rRNA

373 bp

134–137 bp

580 bp

251 bp

398 bp

H. hepaticus

H. muridarum

H. hepaticus

H. bilis

H. typhlonius

H. rodentium

Candidantus H.suis

H.felis

H.bizzozeronii

H.salomonis, H.bizzozeronii, H.felis

H. heilmannii

Helicobacter species

Culture, tissues, feces

Culture, feces, tissues

Culture, tissues

Gastric biospy

Oral cavity, biopsy

50oC2m,95oC10m, (95oC15s,60oC60s) × 45

(94oC1m,55oC1m,72oC1m) × 30

95oC5m, (94oC1m,58oC1m,72oC1m) × 3, (94oC1m,60oC1m,72oC1m) × 35, 72oC7m

94oC3m,57oC2m,72oC3m, (94oC30s,57oC30s,72oC1m) × 30, 94oC20s,57oC20s,72oC5m

94oC5m, (94oC30s,57oC30s,72oC30s) × 35, 72oC7m

94oC5m, (94oC30s,57oC30s,72oC30s) × 40, 72oC7m

2001

2005

2004

1998

1998

2007

(Continued)

86

85

82

83

40

71

Helicobacter 193

Primer

Primer Sequence (5’–3’)

∼1,200 bp

GCGGCCGCCCGTCCCGCCG

CCCCCGCCCCGCCGCGGCC GCCTATGACGGGTATCCGGC TCGCCTTCGCAATGAGTTT GCTATGACGGGTATCC

ACTTCACCCCAGTCGCTG GAATAGACGGGGACCC

CGCCCGCCGCGCCCCGCGC CCGTCCCGCCGCCCCCGCC CGGGGTTGCGCTCGTTGC

1F-GC

2R(667–686) C97(1st)

C05(1st) BSF917

BSR1114GC

∼200 bp

∼470 bp

CTCACGACACGAGCTGAC

1R(1018–1035)

∼780 bp

200 bp

Product

1F(254–271)

AATATACGCGCACACCTCTCA TCTGACCAT CACCTGTGCATTTTGGACGA ACCAAGGCWATGACGGGTATC

CGGAGTTAGCCGGTGCTTATT FAM-AACCTTCATCCTCCACGC GGC-TAMRA CTATGACGGGTATCCGGC

R FAM

cdtBR F

cdtBP

16S rRNA (V3 region)

16S rRNA

16S rRNS

16S rRNA

Target Gene

Helicobacter species

Helicobacter species

Helicobacter species

Target Bacteria

Feces, tissues

Feces

Gastric biospy, feces

Samples

94oC10m, (94oC30s,50oC30s,72oC30s) × 35, 72oC5m

94oC4m, (94oC1m,50oC1m,72oC2m) × 30, 72oC5m

95oC10m, (94oC30s,55oC30s,72oC30s) × 35, 72oC5m

94oC2m, (94oC30s,55oC30s,72oC30s) × 30, 72oC5m

50oC2m,95oC10m, (95oC15s,60oC1m) × 40

Reactions

1993

2004 1998

2006

2003

2004

Year

48

80 40

81

74

75

Reference

Abbreviations:   cdtB, cytolethal distending toxin B; ure, urease; FAM, 6-carboxyfluorescein; TAMRA, 6-carboxytetramethylrhodamine; LAMP, loop-mediated isothermal amplification; DGGE, denaturing gradient gel electrophoresis; °C, temperature in Celsius; m, minutes; s, seconds; ( ) ×, (reaction condition) and cycle; N=A/T/C/G, B=C/G/T, D=A/G/T, R=A/G, S=G/C, W=A/T, Y=C/T.

PCR-DGGE (nested PCR)

PCR-DGGE (Seminested PCR)

Quantitative PCR

Method

Table 13.6  (Continued)

194 Molecular Detection of Foodborne Pathogens

195

Helicobacter

Helicobacter genus. The set of primers (C97/C05) designed by Fox et al. and the primers adjacent to C97/ C05, which target the 16S rRNA gene, are frequently used.40 Gastric biopsies, feces, and tissues from the liver, gallbladder, intestinal tract, and oral cavity are used as samples. While standard PCR is enough to amplify the 16S rRNA gene when DNA is obtained from cultures and tissues, the amplification of DNA from feces and the oral cavity by standard PCR is usually difficult. In fact, because the amount of Helicobacter is not constant in samples from feces or the oral cavity, nested PCR had been demonstrated to be suitable for the detection of Helicobacter in these cases.71 Prachasilpchai et al. showed that there is no significant difference in the sensitivity of a histological test and of PCR for the detection of Helicobacter spp. in tissues from canine stomachs.79 Gastric Helicobacter. In animals, several Helicobacter species with host specificity have been isolated. For ­example, H. heilmannii—formerly “Gastrospirillum hominis,” H. felis, H. bizzozeronii, H.salomonis, H. bovis, and Candidatus H. suis have been isolated from stomachs. PCR methods targeting these Helicobacter species have been developed. The 16S rRNA gene can be analyzed for the identification of Helicobacter species by using primers designed for the region that is specific to each Helicobacter. However, the region suitable for primers is limited. Consequently, identification by sequencing of the 16S rRNA gene and by PCR-DGGE has been performed using a primer that is designed to recognize a sequence common to Helicobacter species.40,74,80,81 Culture of H. heilmannii (type II) is particularly difficult to analyze biochemically. Therefore, the detection and identification of H. heilmannii have been performed by genetic analysis using PCR.35,82 In this way, the presence of H. heilmannii in humans, cats, dogs, and pigs has been demonstrated.83,84 Furthermore, RFLP analysis of the ureB fragment obtained by PCR may be informative for identification if an appropriate restriction enzyme site for the discrimination of species is present.83 In addition to the 16S rRNA gene, the ureB gene is also available as a target gene for the identification of H. heilmannii, H. felis, H. pylori, and H. bizzozeronii.35 For the detection of H. salomonis, H. bizzozeronii, and H. felis, a tRNA gene is useful as a target gene.82 Multiplex-PCR with combinations of primers for the tRNA, 16S rRNA, and ureB genes makes it possible to identify each of these three Helicobacter species.82 Enterohepatic Helicobacter. Primers have been designed for the detection of Helicobacter species such as H. hepaticus, H. cetorum, H. equorum, H. ganmani, H. rodentium, H. typhlonicus, H. bilis, and H. muridarum in the intestinal tract and liver.80,85–88 Since these Helicobacter species lack urease activity, the major target gene is the 16S rRNA gene. Shames et al. demonstrated that including 3.0 mM MgCl2 in the PCR mixture is better than 1.5 mM MgCl2 for amplifying a clear, single band for the detection of H. hepaticus from mouse feces.88 In the case of H. hepaticus, H. cetorum, H. rodentium, H. typhlonicus, H. bilis, and H. muridarum, the set of primers targeting the 16S rRNA gene for the specific

detection of each Helicobacter species have been used successfully in multiplex-PCR.85 For the case of H. ­equorum, specific primers targeting the 23S rRNA gene have been developed,87 while primers targeting the internal spacer region between the 16S and 23S rRNA genes have been reported for the identification of H. ganmani.80 The cdt gene, which is a pathogenetic gene present in some enterohepatic Helicobacter such as H. hepaticus and H. pullorum, has been reported to be useful for the detection of these Helicobacter species;89 moreover, TaqMan PCR for the cdt gene has been shown to be suitable for the rapid detection of H. hepaticus.86

13.3 Conclusions and Future Perspectives Isolating bacteria is necessary for investigating the microbial characteristics and pathogenesis of Helicobacter species. Molecular methods allow the detection of Helicobacter, but they do not lead to the isolation of viable bacteria. Culturing, which is the gold standard in the detection of Helicobacter species, requires media, culture conditions, and sample types that must be optimized for each Helicobacter species. Several challenges remain in developing appropriate media and methods for culturing Helicobacter species. Previously, the presence of spiral bacteria other than H. pylori was detected histologically.15,90,91 For the detection of these bacteria, sequences of the 16S rRNA genes obtained from PCR are important for identifying bacteria that are difficult to culture, such as H. heilmannii.34,92 Furthermore, PCR-based methods will be useful for the detection and identification of novel Helicobacter discovered in the future. Immunoassay methods, including immunochromatography and ELISA, are easy and rapid procedures, and their use in the diagnosis of H. pylori infection is expected to increase. Nevertheless, these methods are less useful for detecting Helicobacter species other than H. pylori. Immunological detection of these other species requires commercially available antibodies specific for each Helicobacter species. With the amount of genomic information continuing to rise rapidly, however, genes specific to each Helicobacter species can be identified. This will allow the design of primers and probes for use in PCR for the simultaneous detection and subtyping of the bacteria. In H. pylori, differences in pathogenic genes such as the cagA and vacA genes have been reported to cause strain-specific differences in the pathogenicities of peptic ulcer and gastric cancer. Consequently, typing of the cagA gene may be useful for investigating the relationship between H. pylori infection and gastric cancer. Through the detection of mutations that confer antimicrobial resistance, it is possible to predict the resistance phenotype based on DNA sequencing.47 However, DNA sequencing requires complicated procedures. Although RFLP is simpler than DNA sequencing, a restriction enzyme that recognizes the mutation is required. DGGE is used for identifying Helicobacter species based on sequence ­differences among DNA amplicons, such as the amplicon for the 16S rRNA gene.80,81 If these methods prove applicable to

196

the analysis of pathogenetic genes, they may be useful for diagnosing pathogenesis, as well as for indentifying the species responsible. In addition, two new techniques have been developed for SNP analysis: the PCR Line Probe assay uses PCR and reverse hybridization, and real-time PCR uses probes labeled with fluorescent dyes.93,94 In addition to detecting SNPs, realtime PCR can rapidly detect both the presence and expression level of the target gene.95–97 In this way, real-time PCR can be used to perform phenotypic analyses, such as an analysis of the levels of pathogenesis in bacteria. Microarray analysis can also reveal information about changes in the expression levels of genes in Helicobacter species.98 These and other methods will continue to be developed as effective tools for examining target genes in Helicobacter infections.

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198 74. Al-Soud, W.A. et al. Assessment of PCR-DGGE for the ­identification of diverse Helicobacter species, and application to faecal samples from zoo animals to determine Helicobacter prevalence. J. Med. Microbiol. 52, 765, 2003. 75. Huijsdens, X.W. et al. Detection of Helicobacter species DNA by quantitative PCR in the gastrointestinal tract of healthy individuals and of patients with inflammatory bowel disease. FEMS Immunol. Med. Microbiol. 41, 79, 2004. 76. Higashi, H. et al. Biological activity of the Helicobacter pylori virulence factor CagA is determined by variation in the tyrosine phosphorylation sites. Proc. Natl. Acad. Sci. USA 99, 14428, 2002. 77. Higashi, H. et al. SHP-2 tyrosine phosphatase as an intracellular target of Helicobacter pylori CagA protein. Science 295, 683, 2002. 78. He, Q. et al. Real-time quantitative PCR for detection of Helicobacter pylori. J. Clin. Microbiol. 40, 3720, 2002. 79. Prachasilpchai, W. et al. Diagnosis of Helicobacter spp. infection in canine stomach. J. Vet. Sci. 8, 139, 2007. 80. Nilsson, H.O. et al. High prevalence of Helicobacter species detected in laboratory mouse strains by multiplex ­PCR-denaturing gradient gel electrophoresis and pyrosequencing. J. Clin. Microbiol. 42, 3781, 2004. 81. Zhang, L. et al. Nongastric Helicobacter species detected in the intestinal tract of children. J. Clin. Microbiol. 44, 2276, 2006. 82. Baele, M. et al. Multiplex PCR assay for differentiation of Helicobacter felis, H. bizzozeronii, and H. salomonis. J. Clin. Microbiol. 42, 1115, 2004. 83. Dieterich, C. et al. Presence of multiple Helicobacter heilmannii strains in an individual suffering from ulcers and in his two cats. J. Clin. Microbiol. 36, 1366, 1998. 84. Eaton, K.A. et al. Prevalence and varieties of Helicobacter species in dogs from random sources and pet dogs: animal and public health implications. J. Clin. Microbiol. 34, 3165, 1996. 85. Feng, S. et al. Differential detection of five mouse-infecting Helicobacter species by multiplex PCR. Clin. Diagn. Lab. Immunol. 12, 531, 2005. 86. Ge, Z. et al. Fluorogenic PCR-based quantitative detection of a murine pathogen, Helicobacter hepaticus. J. Clin. Microbiol. 39, 2598, 2001. 87. Moyaert, H. et al. Prevalence of Helicobacter equorum in faecal samples from horses and humans. Vet. Microbiol. 121, 378, 2007. 88. Shames, B. et al. Identification of widespread Helicobacter hepaticus infection in feces in commercial mouse colonies by culture and PCR assay. J. Clin. Microbiol. 33, 2968, 1995. 89. Ceelen, L.M. et al. The cytolethal distending toxin among Helicobacter pullorum strains from human and poultry origin. Vet. Microbiol. 113, 45, 2006. 90. Andersen, L.P. New Helicobacter species in humans. Dig. Dis. 19, 112, 2001. 91. Kusters, J.G., and Kuipers, E.J. Non-pylori Helicobacter infections in humans. Eur. J. Gastroenterol. Hepatol. 10, 239, 1998. 92. Jalava, K. et al. A cultured strain of “Helicobacter heilmannii,” a human gastric pathogen, identified as H. bizzozeronii: evidence for zoonotic potential of Helicobacter. Emerg. Infect. Dis. 7, 1036, 2001. 93. Prokhorenko, I.A. et al. Phenylethynylpyrene-labeled ­oligonucleotide probes for excimer fluorescence SNP analysis of 23S rRNA gene in clarithromycin-resistant Helicobacter pylori strains. Mutat. Res. 599, 144, 2006.

Molecular Detection of Foodborne Pathogens 94. van Doorn, L.J. et al. Accurate prediction of macrolide ­resistance in Helicobacter pylori by a PCR line probe assay for detection of mutations in the 23S rRNA gene: multicenter validation study. Antimicrob. Agents Chemother. 45, 1500, 2001. 95. Lottspeich, C. et al. Evaluation of the novel Helicobacter pylori ClariRes real-time PCR assay for detection and clarithromycin susceptibility testing of H. pylori in stool specimens from symptomatic children. J. Clin. Microbiol. 45, 1718, 2007. 96. McDaniels, A.E. et al. Evaluation of quantitative real time PCR for the measurement of Helicobacter pylori at low concentrations in drinking water. Water Res. 39, 4808, 2005. 97. Schabereiter-Gurtner, C. et al. Novel real-time PCR assay for detection of Helicobacter pylori infection and simultaneous clarithromycin susceptibility testing of stool and biopsy specimens. J. Clin. Microbiol. 42, 4512, 2004. 98. Goto, K. et al. Specific and quantitative detection of PCR products from Clostridium piliforme, Helicobacter bilis, H. hepaticus, and mouse hepatitis virus infected mouse samples using a newly developed electrochemical DNA chip. J. Microbiol. Methods 69, 93, 2007. 99. Liang, S., and Redlinger, T. A protocol for isolating putative Helicobacter pylori from fecal specimens and genotyping using vacA alleles. Helicobacter 8, 561, 2003. 100. Seymour, C. et al. Isolation of Helicobacter strains from wild bird and swine feces. Appl. Environ. Microbiol. 60, 1025, 1994. 101. Butcher, G.P. et al. Use of an ammonia electrode for rapid quantification of Helicobacter pylori urease: its use in the endoscopy room and in the assessment of urease inhibition by bismuth subsalicylate. Digestion 53, 142, 1992. 102. Prince, M.I. et al. The CLO test in the UK: inappropriate reading and missed results. Eur. J. Gastroenterol. Hepatol. 11, 1251, 1999. 103. Munster, D.J. et al. The fate of ingested 14C-urea in the urea breath test for Helicobacter pylori infection. Scand. J. Gastroenterol. 28, 661, 1993. 104. Kosunen, T.U. et al. Diagnostic value of decreasing IgG, IgA, and IgM antibody titres after eradication of Helicobacter pylori. Lancet 339, 893, 1992. 105. Malfertheiner, P. et al. Current concepts in the management of Helicobacter pylori infection: the Maastricht III Consensus Report. Gut 56, 772, 2007. 106. Mitchell, H.M., et al. Antibody to Campylobacter pylori in families of index children with gastrointestinal illness due to C. pylori. Lancet 2, 681, 1987. 107. Ekstrom, A.M. et al. Helicobacter pylori in gastric cancer established by CagA immunoblot as a marker of past infection. Gastroenterology 121, 784, 2001. 108. Kato, M. et al. Clinical usefulness of urine-based enzymelinked immunosorbent assay for detection of antibody to Helicobacter pylori: a collaborative study in nine medical institutions in Japan. Helicobacter 5, 109, 2000. 109. Kato, S. et al. Urine-based enzyme-linked immunosorbent assay for the detection of Helicobacter pylori infection in children. Pediatrics 107, E87, 2001. 110. Patel, P. et al. Salivary antibodies to Helicobacter pylori: screening dyspeptic patients before endoscopy. Lancet 344, 511, 1994. 111. Katsuragi, K. et al. Highly sensitive urine-based ­enzyme-linked immunosorbent assay for detection of antibody to Helicobacter pylori. Helicobacter 3, 289, 1998. 112. Gatta, L. et al. A rapid immunochromatographic assay for Helicobacter pylori in stool before and after treatment. Aliment Pharmacol. Ther. 20, 469, 2004.

Helicobacter 113. Panayotopoulou, E.G. et al. Strategy to characterize the ­number and type of repeating EPIYA phosphorylation motifs in the carboxyl terminus of CagA protein in Helicobacter pylori clinical isolates. J. Clin. Microbiol. 45, 488, 2007. 114. Riley, L.K. et al. Identification of murine helicobacters by PCR and restriction enzyme analyses. J. Clin. Microbiol. 34, 942, 1996. 115. Eaton, K.A. et al. Helicobacter acinonyx sp. nov., isolated from cheetahs with gastritis. Int. J. Syst. Bacteriol. 43, 99, 1993. 116. Fox, J.G. et al. Helicobacter anseris sp. nov. and Helicobacter brantae sp. nov., isolated from feces of resident Canada geese in the greater Boston area. Appl. Environ. Microbiol. 72, 4633, 2006. 117. Patterson, M.M. et al. Helicobacter aurati sp. nov., a ureasepositive Helicobacter species cultured from gastrointestinal tissues of Syrian hamsters. J. Clin. Microbiol. 38, 3722, 2000. 118. Baele, M. et al. Helicobacter baculiformis sp. nov., isolated from feline stomach mucosa. Int. J. Syst. Evol. Microbiol. 58, 357, 2008. 119. Fox, J.G. et al. Helicobacter bilis sp. nov., a novel Helicobacter species isolated from bile, livers, and intestines of aged, inbred mice. J. Clin. Microbiol. 33, 445, 1995. 120. Hanninen, M.L. et al. Culture and characteristics of Helico­ bacter bizzozeronii, a new canine gastric Helicobacter sp. Int. J. Syst. Bacteriol. 46, 160, 1996. 121. Fox, J.G. et al. Helicobacter canadensis sp. nov. isolated from humans with diarrhea as an example of an emerging pathogen. J. Clin. Microbiol. 38, 2546, 2000. 122. Stanley, J. et al. Helicobacter canis sp. nov., a new species from dogs: an integrated study of phenotype and genotype. J. Gen. Microbiol. 139, 2495, 1993. 123. Harper, C.G. et al. Helicobacter cetorum sp. nov., a ureasepositive Helicobacter species isolated from dolphins and whales. J. Clin. Microbiol. 40, 4536, 2002. 124. Franklin, C.L. et al. Isolation of a novel Helicobacter species. Helicobacter cholecystus sp. nov., from the gallbladders of Syrian hamsters with cholangiofibrosis and centrilobular pancreatitis. J. Clin. Microbiol. 34, 2952, 1996. 125. Totten, P.A. et al. Campylobacter cinaedi (sp. nov.) and Campylobacter fennelliae (sp. nov.): two new Campylobacter species associated with enteric disease in homosexual men. J. Infect. Dis. 151, 131, 1985. 126. Van den Bulck, K. et al. Helicobacter cynogastricus sp. nov., isolated from the canine gastric mucosa. Int. J. Syst. Evol. Microbiol. 56, 1559, 2006. 127. Moyaert, H. et al. Helicobacter equorum sp. nov., a ureasenegative Helicobacter species isolated from horse faeces. Int. J. Syst. Evol. Microbiol. 57, 213, 2007. 128. Lee, A. et al. Isolation of a spiral-shaped bacterium from the cat stomach. Infect. Immun. 56, 2843, 1988. 129. Robertson, B.R. et al. Helicobacter ganmani sp. nov., a ureasenegative anaerobe isolated from the intestines of laboratory mice. Int. J. Syst. Evol. Microbiol. 51, 1881, 2001. 130. Fox, J. G. et al. Helicobacter hepaticus sp. nov., a microaerophilic bacterium isolated from livers and intestinal mucosal scrapings from mice. J. Clin. Microbiol. 32, 1238, 1994. 131. Fox, J.G. et al. Helicobacter marmotae sp. nov. isolated from livers of woodchucks and intestines of cats. J. Clin. Microbiol. 40, 2513, 2002. 132. Shen, Z. et al. A novel enterohepatic Helicobacter species ‘Helicobacter mastomyrinus’ isolated from the liver and intestine of rodents. Helicobacter 10, 59, 2005. 133. Simmons, J.H. et al. Helicobacter mesocricetorum sp. nov., A novel Helicobacter isolated from the feces of Syrian hamsters. J. Clin. Microbiol. 38, 1811, 2000.

199 134. Lee, A. et al. Helicobacter muridarum sp. nov., a microaerophilic helical bacterium with a novel ultrastructure isolated from the intestinal mucosa of rodents. Int. J. Syst. Bacteriol. 42, 27, 1992. 135. Fox, J.G. et al. Gastric colonization by Campylobacter pylori subsp. mustelae in ferrets. Infect. Immun. 56, 2994, 1988. 136. Bronsdon, M.A. et al. Helicobacter nemestrinae sp. nov., a spiral bacterium found in the stomach of a pigtailed macaque (Macaca nemestrina). Int. J. Syst. Bacteriol. 41, 148, 1991. 137. Dewhirst, F.E. et al. Phylogeny of Helicobacter isolates from bird and swine feces and description of Helicobacter pametensis sp. nov. Int. J. Syst. Bacteriol. 44, 553, 1994. 138. Shen, Z. et al. Helicobacter rodentium sp. nov., a urease­negative Helicobacter species isolated from laboratory mice. Int. J. Syst. Bacteriol. 47, 627, 1997. 139. Jalava, K. et al. Helicobacter salomonis sp. nov., a canine gastric Helicobacter sp. related to Helicobacter felis and Helicobacter bizzozeronii. Int. J. Syst. Bacteriol. 47, 975, 1997. 140. Mendes, E.N. et al. Helicobacter trogontum sp. nov., isolated from the rat intestine. Int. J. Syst. Bacteriol. 46, 916, 1996. 141. Franklin, C.L. et al. Helicobacter typhlonius sp. nov., a novel murine urease-negative Helicobacter species. J. Clin. Microbiol. 39, 3920, 2001. 142. Alm, R.A. et al. Genomic-sequence comparison of two unrelated isolates of the human gastric pathogen Helicobacter pylori. Nature 397, 176, 1999. 143. Tomb, J. F. et al. The complete genome sequence of the gastric pathogen Helicobacter pylori. Nature 388, 539, 1997. 144. O’Rourke, J.L. et al. Description of ‘Candidatus Helicobacter heilmannii’ based on DNA sequence analysis of 16S rRNA and urease genes. Int. J. Syst. Evol. Microbiol., 54, 2203, 2004. 145. Zambon, C.F. et al. Non-invasive diagnosis of Helicobacter pylori infection: simplified 13C-urea breath test, stool antigen testing, or DNA PCR in human feces in a clinical laboratory setting? Clin. Biochem. 37, 261, 2004. 146. van Doorn, L.J. et al. Typing of Helicobacter pylori vacA gene and detection of cagA gene by PCR and reverse hybridization. J. Clin. Microbiol. 36, 1271, 1998. 147. Roosendaal, R. et al. Slaughter pigs are commonly infected by closely related but distinct gastric ulcerative lesion-inducing gastrospirilla. J. Clin. Microbiol., 38, 2661, 2000. 148. Welsh, J., and McClelland, M. Genomic fingerprints produced by PCR with consensus tRNA gene primers. Nucleic Acids Res. 19, 861, 1991. 149. Shen, Z. et al. Development of a PCR-restriction fragment length polymorphism assay using the nucleotide sequence of the Helicobacter hepaticus urease structural genes ureAB. J. Clin. Microbiol. 36, 2447, 1998. 150. Ceelen, L. et al. Prevalence of Helicobacter pullorum among patients with gastrointestinal disease and clinically healthy persons. J. Clin. Microbiol. 43, 2984, 2005. 151. Harper, C.G. et al. Comparison of diagnostic techniques for Helicobacter cetorum infection in wild Atlantic bottlenose dolphins (Tursiops truncatus). J. Clin. Microbiol. 41, 2842, 2003. 152. Maeda, S. et al. Helicobacter pylori specific nested PCR assay for the detection of 23S rRNA mutation associated with clarithromycin resistance. Gut 43, 317, 1998. 153. Clayton, C.L. et al. Sensitive detection of Helicobacter pylori by using polymerase chain reaction. J. Clin. Microbiol. 30, 192, 1992. 154. Contreras, M. et al. Detection of Helicobacter-like DNA in the gastric mucosa of Thoroughbred horses. Lett. Appl. Microbiol. 45, 553, 2007.

200 155. Rugge, M. et al. Patients younger than 40 years with gastric carcinoma: Helicobacter pylori genotype and associated gastritis phenotype. Cancer 85, 2506, 1999. 156. Kawamata, O. et al. Nested-polymerase chain reaction for the detection of Helicobacter pylori infection with novel primers designed by sequence analysis of urease A gene in clinically isolated bacterial strains. Biochem. Biophys. Res. Commun. 219, 266, 1996.

Molecular Detection of Foodborne Pathogens 157. Beckwith, C.S. et al., Fecal PCR assay for diagnosis of Helicobacter infection in laboratory rodents. J. Clin. Microbiol. 35, 1620, 1997. 158. Neubauer, C., and Hess, M. Detection and identification of food-borne pathogens of the genera Campylobacter, Arcobacter, and Helicobacter by multiplex PCR in poultry and poultry products. J. Vet. Med. B Infect. Dis. Vet. Public Health 53, 376, 2006.

14 Kocuria

Edoardo Carretto and Daniela Barbarini Fondazione IRCCS Policlinico San Matteo

Contents 14.1 Introduction.................................................................................................................................................................... 201 14.1.1 Classification of Kocuria species...................................................................................................................... 201 14.1.2 Biology, Pathogenesis, and Medical Importance.............................................................................................. 201 14.1.3 Kocuria and Food............................................................................................................................................. 202 14.1.4 Current Diagnostic Methods for Kocuria spp.................................................................................................. 202 14.2 Methods.......................................................................................................................................................................... 203 14.2.1 Automated Ribotyping...................................................................................................................................... 203 14.2.2 PCR Amplification of 16S rDNA Region......................................................................................................... 204 14.2.3 PCR Amplification of the 16S±23S rDNA Intergenic Region......................................................................... 204 14.3 Conclusions and Future Perspectives............................................................................................................................. 204 References.................................................................................................................................................................................. 205

14.1  Introduction The genus Kocuria belongs to the family Micrococcaceae, order Actinomycetales. It was created in 1995 on the basis of the phylogenetic and chemotaxonomic dissection of the genus Micrococcus.1 These microrganisms are Gram-positive cocci, catalase positive, coagulase negative, strictly aerobic, occurring often in tetrads or irregular clusters, and usually grow on simple media. In humans, they may colonize the skin, mucosae and oropharynx; they are also isolated from a great variety of animal sources and from other environments. Considering its relatively small genome size among actinomycetes, the existence of different Kocuria species suggests their capacity to adapt and prosper in their own ecological niche.2

14.1.1  Classification of Kocuria species At it stands, the genus Kocuria consists of 12 valid species: Kocuria rosea (the type species),1 K. varians,1 K. kristinae,1 K. palustris,3 K. rhizophila3 (whose complete genome was recently sequenced2), K. polaris,4 K. marina,5 K. carniphila,6 K. aegyptia,7 K. himachalensis,8 K. flava,9 and K. turfanensis.9 These species were recovered from different environments such as saline, alkaline desert (K. aegyptia), meat (K. carniphila), marine sediment (K. marina), the rhizoplane of narrowleaved cattail (K. palustris, K. rhizophila), an Antarctic cyanobacterial mat sample (K. polaris) and from plates used to detect cultivable bacteria from the air (K. flava and K. turfanensis).9 In general, the members of this genus are Gram-positive, aerobic, coccoid, nonencapsulated, nonhalophilic, and nonendospore-forming, with the presence of the fatty acid anteiso-C15:0 and MK-7(H2) and MK-8(H2) as the major

menaquinones. On Columbia blood agar, Kocuria kristinae (the species most frequently isolated in human infections) typically forms cream colonies. It is nonhemolytic, catalase positive, strictly aerobic, nonmotile, coagulase negative, and unable to reduce nitrate to nitrites and to hydrolyze gelatin, arginine and esculine. K. kristinae produces acid from threalose, glucose, fructose, mannose, glycerol, saccharose but not from mannitol, raffinose, arabinose, lactose, and ribose. The morphological characteristics of the other Kocuria species are variable and are fully illustrated in the first description of a species.1–9

14.1.2 Biology, Pathogenesis, and Medical Importance Up to 1995, members of the genus Kocuria have been regarded as part of the genus Micrococcus in the family Micrococcaceae. Micrococcus strains have been isolated from different clinical specimens, although their clinical relevance is often questionable. From a clinical point of view, Micrococcus luteus is the microrganism that was most ­frequently associated with diseases in humans. There are reports of bacteremia,10,11 pneumonia,12 intracranial abscesses,13 septic arthritis,14 and meningitis,15 either in immunosuppressed or immunocompetent hosts. Different reports suggest also that Micrococcus and related organisms can be involved in peritonitis in patients undergoing ambulatory peritoneal dialysis,16 endocarditis,17 or infection of cerebrospinal fluid shunts.18 Some authors found a significant association between Micrococcus catheter-related bacteremias and patients receiving epoprostenol by continuous intravenous infusion.19 After the rearrangement of the family Micrococcaceae in 1995, generating the new genus Kocuria, some reports 201

202

emphasized that two members of this genus (K. kristinae and K. rosea) were associated to human diseases. Since these microrganisms may colonize the skin, mucosae and oropharynx, it is not surprising that both species were involved in catheter-related bacteremias,20–22 and K. kristinae was also associated to a case of acute cholecystitis.23 It is possible that the true incidence of the bacteremia due to Kocuria spp. may be underestimated due to the following reasons: (i) the correct identification of micrococci isolated from clinical specimens is often cumbersome, since standard biochemical analysis does not work well on these microrganisms, which demonstrate phenotypic variability; (ii) the clinical interpretation of positive results may be misleading; (iii) these microorganisms are hastily discarded as contaminants. The repeated isolation from different blood cultures in the absence of other microorganisms, together with the isolation from a catheter, strongly suggests that Kocuria spp. may cause catheter-related bacteremia. Although this microorganism is intrinsically low-virulent and its presence in the bloodstream is often the expression of a transient bacteremia, the clinicians should not underestimate the repeated isolation of a Kocuria strain from blood cultures, considering the clinical signs of a sepsis syndrome resulting from catheterrelated infection.20 There are relatively few reports on literature about the drug susceptibilities of Kocuria spp. All the strains causing infections were susceptible to oxacillin, first generation cephalosporines, amoxicillin-clavulanate (these β-lactams can be considered as the drug of choice in case of infections due to Kocuria spp.), tetracyclines, glycopeptides, with a different degree of susceptibilities to penicillin and macrolides.20,21,23 To date, the link between Kocuria spp. and foodborne diseases has not been established; and these microorganisms have not been investigated at all in the routine panel of microbiological analysis relating to diarrheal illness. Given that Kocuria species isolated in humans (K. kristinae, K. rosea, K. varians) show the following biological characteristics: (i) colonizing microorganisms of the skin, mucosae and oropharynx, without any ability to cause systemic diseases (with the exception of foreign-body associated infections and a case of cholecystitis) or localized diseases (skin or soft tissues infections were never reported) and (ii) the lack of evidence of genes with particular virulence and with toxigenic activity, it is postulated that these microorganisms have no ability to cause intestinal diseases in humans.

14.1.3  Kocuria and Food Species belonging to the genera Pediococcus, Lactobacillus, Kocuria, and Staphylococcus are often used in the manufacture of fermented sausages as starter cultures. The most important roles of coagulase-negative staphylococci (CNS) and Kocuria spp. are: (i). the reduction of nitrate to nitrite, to promote the desired red color development and stability; (ii) peroxide decomposition and therefore, prevention of rancidity; and (iii) nitrite permanence with consequent maintenance of hygienic safety. Furthermore, the lipolytic and proteolytic activity of these bacteria may contribute to the

Molecular Detection of Foodborne Pathogens

flavor of fermented meat products by formation of esters and other aromatic compounds from amino acids. Mixed cultures of lactic acid bacteria and Staphylococcus xylosus, S. carnosus, S. saprophyticus, and K. varians are increasingly being used to standardize the quality of fermented sausages.24,25 Interestingly, certain strains are able to inhibit spoilage or pathogenic bacteria; in particular, the potential antilisterial activity of S. xylosus, S. carnosus, and Kocuria varians was investigated by a number of authors.26,27 A recent paper examined the in vitro and in vivo interactions occurring between S. xylosus and K. varians in the ripening of traditional fermented meat products of southern Italy. This study pointed out that S. xylosus strains produced unidentified compounds capable of positively or negatively affecting the growth of K. varians; and that a high stimulatory interaction assessed in vitro by using mixtures of K. varians K4 and certain S. xylosus strains, resulted in a strongly proteolytic activity on sarcoplasmic proteins, whereas a moderate stimulatory interaction, found between K. varians K4 and other S. xylosus strains, corresponded to a lower proteolytic activity.28

14.1.4 Current Diagnostic Methods for Kocuria spp. Usually, the strains of Kocuria spp. having medical interest grow well on simple nonselective media such as Columbia blood agar. Using a hierarchical approach to identify Kocuria spp., the first methods that can be used are phenotypic. Among the different phenotypic methods commercial systems, such as the API 20 Staph or the ID32 Staph ATB system (both from BioMérieux, Marcy l’Etoile, France) has been proven useful. The API 20 Staph system consists of a strip containing 20 dehydrated test substrates in single microtubes. These substrates are reconstituted by adding to each tube an aliquot of the medium, provided by the manufacturer, that has been inoculated with the strain to be studied. After a 24 h incubation at 35°C the reaction results were read and compared to a reference identification database provided by the manufacturer.29 The ID32 Staph ATB system is another standardized system consisting of 32 cupules, 26 of which are used as test cupules containing dehydrated test substrates. These substrates are reconstituted by adding to each tube an aliquot of the medium, provided by the manufacturer, that has been inoculated with the strain to be studied. After a 24 h incubation at 36°C the reaction results can be read either using the ATB™ Expression™ or miniAPI® instruments, or visually. The obtained results should be compared to a reference identification database provided by the manufacturer.30 Various automated identification and susceptibility test systems are currently on the market, among them Vitek 2 (BioMérieux, Marcy l’Etoile, France) and the BD Phoenix system (Becton Dickinson Diagnostic Systems, Sparks, MD). In a previous paper, Ben-Ami reported the possibility that Vitek 2 could misidentify S. epidermidis strains as Kocuria spp.31 However, quite recently, the Vitek 2 Grampositive identification card has been redesigned to increase

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Kocuria

the accuracy in the identification of Gram-positive cocci; some Author pointed out that this new card seems to work on Kocuria strains better than the previous one.32 Finally, it has to be emphasized that there are evidences that, in general, manual systems work better on CNS compared with the automated systems.33 Unfortunately, use of phenotypic procedures to identify CNS can be problematic. Commercial identification kits or automated systems often provide unreliable results for most CNS species, in particular non S. epidermidis isolates. This is attributed mainly to the variable nature of diagnostic reactions applied to the species and to the subjectivity in interpretation.34 Misidentification of Kocuria species has also been described for the same reasons.31 Thus, the correct identification of an isolate phenotypically identified as Kocuria species should be confirmed using molecular techniques. The sequencing of the region encoding for the 16S rDNA was proven useful to correctly identify these bacteria.20,23 Ribotyping was proven very useful in the study of staphylococci34 both for molecular epidemiology purposes and for identification and taxonomic purposes. High quality databases of ribotyping profiles that include representative strains of nearly all described species and subspecies of CNS have been constructed and are useful for identification of unknown isolates.35 The RiboPrinter® Microbial Characterization System (DuPont-Qualicon, Wilmington, DE) is an automated instrument able to perform the ribotyping of bacterial isolates. After the generation of an electronic ribotype fingerprint, the instrument software compares it automatically to the reference DuPont identification databases. In the past, we demonstrated the usefulness of the RiboPrinter® in identifying CNS.34 In our experiments we did not include any Kocuria strains, since at that time the DuPont libraries that the instrument uses to identify the strains did not include Kocuria profiles. However, the most updated versions of the DuPont databases (version 6.8 for the EcoRI library and version 1.9 for the PvuII library) contain 26 different profiles of four different species of Kocuria: K. kristinae (five profiles), K. rosea (seven profiles), K. rhizophila (two profiles), and K. varians (11 profiles). In our experience this instrument demonstrated very useful to correctly identify Kocuria strains (unpublished data) and could be used as a first line molecular approach to confirm the putative phenotypical identification of these microrganisms, instead of the 16S rDNA sequencing. Since the food industry needs to check rapidly the presence and/or to monitor during fermentation the inoculated starter cultures, and the reliability of the identification of strains used as starter can have legal importance, efforts were made in developing molecular systems able to provide a correct identification for CNS. The gradient gel electrophoresis (DGGE) analysis of PCR amplified 16S ribosomal sequences has been applied for identification of Micrococcaceae involved in sausage fermentation. The results obtained showed that strains of S. xylosus, S. cohnii, S. intermedius, S. carnosus, K. varians e K. kristinae could be reliably identified on the basis of the analysis of the DGGE mobility of the 16S rDNA V3 region.36 More recently, a polyphasic approach combining the16S–23S

rDNA intergenic spacer region (ISR)-polymerase chain reaction (PCR) and the 16S rDNA V3 region PCR-DGGE techniques has been demonstrated very useful in determining the staphylococcal population of a fermented food product.37 Rossi et al. used a random amplified polymorphic DNA technique based on the rDNA 16S±23S spacer region amplification to identify strains belonging to S. xylosus, S. simulans, S. carnosus or closely related species and of Kocuria species in the sausage microflora, confirming the usefulness of this technique in differentiating among these bacterial species.25 To our knowledge, a number of PCR amplicon sequencingbased methods for identification of CNS have been reported, i.e., targeting the 16S rDNA gene,38 the manganese-dependent superoxide dismutase (sodA) gene,39 the glyceraldehyde3-phosphate dehydrogenase-encoding gene (gap gene).40 However none of the above molecular techniques have been evaluated (and therefore have been proven useful) to identify Kocuria spp.

14.2  Methods To confirm the appropriateness of the identification of Kocuria spp. it is often necessary to use molecular methods. Here we describe some quick protocol to identify these strains: (i) using the RiboPrinter® Microbial Characterization System; (ii) amplifying the 16S rDNA region; and (iii) amplifying the 16S±23S rDNA intergenic region.

14.2.1  Automated Ribotyping Isolated bacterial colonies of the putative Kocuria strains, grown in different culture media, should be subcultured in brain heart infusion agar. Two different experiments should be performed, since the isolates are better analyzed using either EcoRI or PvuII as restriction enzyme. After an overnight growth at 37°C on this medium, isolated colonies are collected using a plastic stick and suspended into the supplied sample buffer. The correct inoculum should be very heavy (four colonies in 40 µl of buffer). The bacterial suspension is then treated for 10 min at 80°C, to reduce viability and to inactivate nucleases. After addition of 5 µl of two lytic enzymes (supplied by the manufacturer) to 30 µl of the bacterial suspensions, samples can be loaded into the RiboPrinter® system together with the chosen restriction enzyme. From now on, all the process is fully automated and performed by the instrument. After the enzyme digestions, the protocol includes separation of restriction fragments by agarose gel electrophoresis, blotting onto a Nylon membrane, and hybridization with a chemiluminescently labeled rDNA probe corresponding to the Escherichia coli rrnB operon. The patterns obtained are then read by a Charge Coupled Device camera, and restriction patterns are automatically analyzed, producing normalized digital fingerprints that are stored in the computer internal database. After generation of the electronic ribotype fingerprint, the instrument software compares it automatically to the reference DuPont identification database. To date, the most updated versions of the DuPont databases

204

(version 6.8 for the EcoRI library and version 1.9 for the PvuII library) contain 26 different profiles of four different species of Kocuria: K. kristinae (three EcoRI profiles, two PvuII profiles), K. rosea (four EcoRI profiles, three PvuII profiles), K. rhizophila (two EcoRI profiles), and K. varians (two EcoRI profiles, ten PvuII profiles). For identification, a threshold value of 85% overall similarity (as determined by using an algorithm similar to the Pearson correlation coefficient in the range of molecular sizes from 1 to 48 kb) between the test fingerprint and the database reference fingerprint is used.

14.2.2  PCR Amplification of 16S rDNA Region Isolated bacterial colonies of the putative Kocuria strains, grown in different culture media, should be subcultured in nonselective media, such as brain heart infusion agar or Columbia blood agar. After the extraction of genomic DNA using conventional methods, the amplification of the 16S rDNA can be achieved using universal primers. Two of the most commonly used primers for bacterial 16S rDNA genes are 27F (5′-AGAGTTTGATCCTGGCTCAG-3′) spanning positions eight to 27 in Escherichia coli rDNA coordinates, and 1492R (5′-TACCTTGTTACGACTT-3′), commonly spanning positions 1492–1507, though longer versions are sometimes used, which amplify nearly the entire length of the gene.20,41–43 As an example, PCR mixtures can contain 50 mM KCl, 10 mM Tris (pH 8.3), 1.5 mM MgCl2, 2 mM each of the four deoxynucleoside triphosphates, 1 µM (each) oligonucleotide primer, 2.5 U of Taq DNA polymerase, and 1 pg to 100 ng of template DNA in a total reaction volume of 100 µl. Amplifications are carried out on a thermal cycler with a previous initial denaturation at 94°C for 5 min, followed by 25 cycles of 94°C for 60 sec, 50°C for 30 sec, and 72°C for 2 min, with a final 5 min extension step at 72°C. The amplicons can be kept at 4°C until purified. The PCR products can be sequenced using the ABI Big Dye Terminator technology with the same primers. The nucleotide sequences can be compared to GenBank contents using the BLAST program. Recently, we successfully used this approach to confirm the identification of a strain which caused repeated episodes of bacteremia; the nucleotide sequence, analyzed with BLAST program, revealed a higher homology (99.8%) with K. kristinae (AF323746). The sequence has been deposited in the GenBank/EMBL data library under accession number AJ316579.20 Commercial 16S rDNA sequencing kits are nowadays available to identify bacteria and some Authors point out that this molecular approach can be useful for strains expressing ambiguous biochemical profiles.44 However, to our knowledge these commercial systems have not been evaluated for the identification of Kocuria strains.

14.2.3 PCR Amplification of the 16S±23S rDNA Intergenic Region Isolated bacterial colonies of the putative Kocuria strains, grown in different culture media, should be subcultured in nonselective media, such as brain heart infusion agar

Molecular Detection of Foodborne Pathogens

or Columbia blood agar. After the extraction of genomic DNA using conventional methods, the amplification of the 16S±23S rDNA intergenic region can be achieved using universal primers.45 In a recent paper, Rossi et al. used primers OPL-01 (5′-GGCATGACCT-3′), OPL-02 (5′-TGGGCGTCAA-3′), OPL-05 (5′-ACGCAGGCAC-3′) and Hpy 1 (5′-CCGCAGCCAA-3′). PCR mixtures contained Taq buffer, 1.5 mM MgCl2, 2 mM each of the four deoxynucleoside triphosphates, 2.5 µM (each) of oligonucleotide primers, 2.5 U of Taq DNA polymerase, and 50 ng of template DNA in a total reaction volume of 50 µl. The amplification programme comprises initial denaturation at 94°C for 4 min, 45 cycles of denaturation at 94°C for 30 sec, annealing at 35°C for 30 sec, and extension at 72°C for 1 min, with a final extension at 72°C for 5 min. The electrophoresis on agarose gel of the PCR products allow to detect a single band at 500 bp for K. varians, at 550 bp for K. kristinae and a single band at 650 bp for K. rosea.25

14.3 Conclusions and Future Perspectives The genus Kocuria consists of a group of Gram-positive, catalase positive, coagulase negative, strictly aerobic cocci that have been long considered as part of the genus Micrococcus in the family Micrococcaceae. Although Micrococcus species such as M. luteus have been isolated from a variety of clinical specimens and implicated in several human diseases (e.g., catheter-related bacteremia, pneumonia, intracranial abscesses, septic arthritis, meningitis, peritonitis and endocarditis, etc.), there is a conspicuous lack of evidence concerning the ability of Kocuria species to induce intestinal (or foodborne) diseases in humans, despite the fact that Kocuria spp. are found in various animal, food and environmental specimens, and have the capacity to colonize the skin, mucosae, and oropharynx in human host; and that K. kristinae and K. rosea have been also associated with catheter-related bacteremia and/or acute cholecystitis. Given the close relationship between Kocuria and Micrococcus genera, and the relative short existence of Kocuria as an independent genus, it is possible that the role of Kocuria spp. in the causation of bacteremia and other human diseases may have been overlooked. Certainly, the current difficulty to correctly identify micrococcal isolates from clinical specimens by conventional procedures and the tendency to dispose the isolated micrococci as contaminants without detailed analysis may have contributed to the dearth of data on human diseases due to Kocuria spp. Considering that Kocuria species, along with Pediococcus, Lactobacillus, and Staphylococcus, are commonly applied in food industry as starter cultures, it is important to further verify the potential involvement of these bacteria in the initiation of disease process or exacerbation of the underlying diseases in mammalian hosts. The development and application of molecular tests for rapid, sensitive, and specific identification and detection of Kocuria species are not only vital to pinpoint the roles of these bacteria in disease initiation and/or exacerbation, but also provide a

Kocuria

differential diagnosis that will confirm or rule out the potential involvement of these bacteria in foodborne disease outbreaks. Future refinement of sample preparation procedures and adoption of real-time detection formats will further enhance the utility and performance of molecular tests for direct identification and differentiation of Kocuria spp. from clinical, food and environmental specimens.

References





1. Stackebrandt, E. et al. Taxonomic dissection of the genus Micrococcus: Kocuria gen. nov., Nesterenkonia gen. nov., Kytococcus gen. nov., Dermacoccus gen. nov., and Micrococcus Cohn 1872 gen. emend. Int. J. Syst. Bacteriol., 45, 682, 1995. 2. Takarada, H. et al. Complete genome sequence of the soil actinomycete Kocuria rhizophila. J. Bacteriol., 190, 4139, 2008. 3. Kovacs, G. et al. Kocuria palustris sp. nov. and Kocuria rhizophila sp. nov., isolated from the rhizoplane of the narrow-leaved cattail (Typha angustifolia). Int. J. Syst. Bacteriol., 49, 167, 1990. 4. Reddy, G.S. et al. Kocuria polaris sp. nov., an orange-pigmented psychrophilic bacterium isolated from an Antarctic cyanobacterial mat sample. Int. J. Syst. Evol. Microbiol., 53, 183, 2003. 5. Kim, S.B. et al. Kocuria marina sp. nov., a novel actinobacterium isolated from marine sediment. Int. J. Syst. Evol. Microbiol., 54, 1617, 2004. 6. Tvrzova, L. et al. Reclassification of strain CCM 132, previously classified as Kocuria varians, as Kocuria carniphila sp. Nov. Int. J. Syst. Evol. Microbiol., 55, 139, 2005. 7. Li, W.J. et al. Kocuria aegyptia sp. nov., a novel actinobacterium isolated from a saline, alkaline desert soil in Egypt. Int. J. Syst. Evol. Microbiol., 56, 733, 2006. 8. Mayilraj, S. et al. Kocuria himachalensis sp. nov., an actinobacterium isolated from the Indian Himalayas. Int. J. Syst. Evol. Microbiol., 56, 1971, 2006. 9. Zhou, G. et al. Kocuria flava sp. nov. and Kocuria turfanensis sp. nov., airborne actinobacteria isolated from Xinjiang, China. Int. J. Syst. Evol. Microbiol., 58, 1304, 2008. 10. Peces, R. et al. Relapsing bacteraemia due to Micrococcus luteus in a haemodialysis patient with a Perm-Cath catheter. Nephrol. Dial. Transplant., 12, 2428, 1997. 11. von Eiff, C. et al. Micrococcus luteus as a cause of recurrent bacteremia. Pediatr. Infect. Dis. J., 15, 711, 1996. 12. Souhami, L. et al. Micrococcus luteus pneumonia: a case report and review of the literature. Med. Pediatr. Oncol., 7, 309, 1979. 13. Selladurai, B.M. et al. Intracranial suppuration caused by Micrococcus luteus. Br. J. Neurosurg., 7, 205, 1993. 14. Wharton, M. et al. Septic arthritis due to Micrococcus luteus. J. Rheumatol., 13, 659, 1986. 15. Fosse, T. et al. Meningitis due to Micrococcus luteus. Infection, 13, 280, 1985. 16. Spencer, R.C. Infections in continuous ambulatory peritoneal dialysis. J. Med. Microbiol., 27, 1, 1988. 17. Richardson, J.F., Marples, R.R. and de Saxe, M.J. Characters of coagulase-negative staphylococci and micrococci from cases of endocarditis. J. Hosp. Infect., 5, 164, 1984. 18. Shapiro, S. et al. Origin of organisms infecting ventricular shunts. Neurosurg. 22, 868, 1988. 19. Yap, R.L. and Mermel, L.A. Micrococcus infection in patients receiving epoprostenol by continuous infusion. Eur. J. Clin. Microbiol. Infect. Dis., 22, 704, 2003.

205 20. Basaglia, G. et al. Catheter-related bacteremia due to Kocuria kristinae in a patient with ovarian cancer. J. Clin. Microbiol., 40, 311, 2002. 21. Altuntas, F. et al. Catheter-related bacteremia due to Kocuria rosea in a patient undergoing peripheral blood stem cell transplantation. BMC Infect. Dis., 4, 62, 2004. 22. Martinaud, C. et al. Bacteremia caused by Kocuria kristinae in a patient with acute leukaemia. Med. Mal. Infect., 38, 165, 2008. 23. Ma, E.S. et al. Kocuria kristinae infection associated with acute cholecystitis. BMC Infect. Dis., 5, 60, 2005. 24. Martin, B. et al. Molecular, technological and safety characterization of Gram-positive catalase-positive cocci from slightly fermented sausages. Int. J. Food Microbiol., 107, 148, 2006. 25. Rossi, F. et al. Identification by 16S-23S rDNA intergenic region amplification, genotypic and phenotypic clustering of Staphylococcus xylosus strains from dry sausages. J. Appl. Microbiol., 90, 365, 2001. 26. Ryser, E.T. et al. Isolation and identification of cheese-smear bacteria inhibitory to Listeria spp. Int. J. Food Microbiol., 21, 237, 1994. 27. Papamanoli, E. et al. Characterization of Micrococcaceae isolated from dry fermented sausage. Food Microbiol., 19, 441, 2002. 28. Tremonte, P. et al. Interactions between strains of Staphylococcus xylosus and Kocuria varians isolated from fermented meats. J. Appl. Microbiol., 103, 743, 2007. 29. Gemmell, C.G. and Dawson, J.E. Identification of coagulasenegative staphylococci with the API staph system. J. Clin. Microbiol., 16, 874, 1982. 30. Chesneau, O. et al. Usefulness of the ID32 staph system and a method based on rRNA gene restriction site polymorphism analysis for species and subspecies identification of staphylococcal clinical isolates. J. Clin. Microbiol., 30, 2346, 1992. 31. Ben-Ami, R. et al. Erroneous reporting of coagulase-negative Staphylococci as Kocuria spp. by the Vitek 2 system. J. Clin. Microbiol., 43, 1448, 2005. 32. Boudewijns, M. et al. Vitek 2 automated identification system and Kocuria kristinae. J. Clin. Microbiol., 43, 5832, 2005. 33. Layer, F. et al. Comparative study using various methods for identification of Staphylococcus species in clinical specimens. J. Clin. Microbiol., 44, 2824, 2006. 34. Carretto, E. et al. Identification of coagulase-negative staphylococci other than Staphylococcus epidermidis by automated ribotyping. Clin. Microbiol. Infect., 11, 177, 2005. 35. Chesneau, O. et al. The value of rRNA gene restriction site polymorphism analysis for delineating taxa in the genus Staphylococcus. Int. J. Syst. Evol. Microbiol., 50, 689, 2000. 36. Cocolin, L. et al. A novel polymerase chain reaction (PCR)denaturing gradient gel electrophoresis (DGGE) for the identification of Micrococcaceae strain involved in meat fermentations. Its application to naturally fermented Italian sausages. Meat Sci., 57, 59, 2001. 37. Blaiotta, G. et al. Combining denaturing gradient gel electrophoresis of 16S rDNA V3 region and 16S-23S rDNA spacer region polymorphism analyses for the identification of staphylococci from Italian fermented sausages. Syst. Appl. Microbiol., 26, 423, 2003. 38. Becker, K. et al. Development and evaluation of a ­quality-controlled ribosomal sequence database for 16S ribosomal DNA-based identification of Staphylococcus species. J. Clin. Microbiol., 42, 4988, 2004.

206 39. Blaiotta, G., Casaburi, A. and Villani, F. Identification and differentiation of Staphylococcus carnosus and Staphylococcus simulans by species-specific PCR assays of sodA genes. Syst. Appl. Microbiol., 28, 519, 2005. 40. Yugueros, J. et al. Glyceraldehyde-3-phosphate dehydrogenase-encoding gene as a useful taxonomic tool for Staphylococcus spp. J. Clin. Microbiol., 38, 4351, 2000. 41. Lane, D.J. et al. Rapid determination of 16S ribosomal RNA sequences for phylogenetic analyses. Proc. Natl. Acad. Sci. USA, 82, 6955, 1985. 42. Weisburg, W.G. et al. 16S ribosomal DNA amplification for phylogenetic study. J. Bacteriol., 173, 697, 1991.

Molecular Detection of Foodborne Pathogens 43. Wilson, K.H., Blitchington, R.B. and Greene, R.C. Amplification of bacterial 16S ribosomal DNA with polymerase chain reaction. J. Clin. Microbiol., 28, 1942, 1990. 44. Woo, P.C. et al. Usefulness of the MicroSeq 500 16S ribosomal DNA-based bacterial identification system for identification of clinically significant bacterial isolates with ambiguous biochemical profiles. J. Clin. Microbiol., 41, 1996, 2003. 45. Jensen, M.A., Webster, J.A. and Straus, N. Rapid identification of bacteria on the basis of polymerase chain reactionamplified ribosomal DNA spacer polymorphisms. Appl. Environ. Microbiol., 59, 945, 1993.

15 Listeria

Dongyou Liu

Mississippi State University

Hans-Jürgen Busse

University of Veterinary Medicine Vienna

Contents 15.1 Introduction.................................................................................................................................................................... 207 15.1.1 Classification, Morphology, and Biology.......................................................................................................... 207 15.1.2 Epidemiology ................................................................................................................................................... 208 15.1.3 Clinical Features . ............................................................................................................................................ 209 15.1.4 Pathogenesis.......................................................................................................................................................210 15.1.5 Genomics...........................................................................................................................................................210 15.1.6 Diagnosis........................................................................................................................................................... 212 15.1.6.1 Conventian Techniques.................................................................................................................... 212 15.1.6.2 Molecular Techniques...................................................................................................................... 213 15.2 Methods.......................................................................................................................................................................... 213 15.2.1 Sample Preparation .......................................................................................................................................... 213 15.2.2 Detection Procedures.........................................................................................................................................214 15.2.2.1 Multipex PCR Identification of Listeria Species..............................................................................214 15.2.2.2 Multiplex PCR Differentiation of L. monocytogenes Serogroups . ................................................ 215 15.2.2.3 Multipex PCR Determination of L. monocytogenes Virulence ..................................................... 215 15.3 Conclusions and Future Perspectives..............................................................................................................................217 References...................................................................................................................................................................................217

15.1  Introduction 15.1.1  Classification, Morphology, and Biology The genus Listeria encompasses a group of closely related, Gram-positive, nonspore-forming, rod-shaped bacteria of 0.4–0.5 µm×1–1.5 µm in size and a relatively low G+C content ranging from 36 to 39%.1–3 This latter feature in combination with Gram staining behavior implies the relatedness of Listeria to the Firmicutes (Gram-positives with low G+C content) like Bacillus, Brochothrix, Clostridium, Enterococcus, Streptococcus, and Staphylococcus. Taxonomically, the genus Listeria comprises so far six species: L. monocytogenes, L. ivanovii, L. seeligeri, L. innocua, L. welshimeri, and L. grayi, with L. ivanovii being further divided into two subspecies (L. ivanovii subsp. londoniensis and L. ivanovii subsp. ivanovii). Given their ability to endure arduous external conditions including extreme pH, temperature, and salt ranges,4,5 Listeria species are abundantly present in a diversity of environments (e.g., soil, water, effluents, and foods). Whereas L. monocytogenes is a facultative intracellular pathogen of both humans and animals, L. ivanovii (previously known as L. monocytogenes serotype 5) mainly infects ungulated

animals (e.g., sheep and cattle),6 and the other four species are essentially saprophytes that have adapted for free-living in soil and decaying vegetation.7 Using serological techniques that detect interactions between Listeria somatic (O)/flagellar (H) antigens and their corresponding antisera, Listeria can be subdivided into at least 15 serovars, with L. monocytogenes consisting of serovars 1/2a, 1/2b, 1/2c, 3a, 3b, 3c, 4a, 4ab, 4b, 4c, 4d, 4e, and 7; L. ivanovii of serovar 5; L. innocua of serovars 4ab, 6a, and 6b; nd L. welshimeri of serovars 6a and 6b; L. seeligeri of 1/2b, 4c, 4d, and 6b; and L. grayi of serovar Grayi (Table 15.1).8 Upon various genetic typing analyses, L. monocytogenes is grouped into three genetic lineages, with lineage I comprising serovars 1/2b, 3b, 4b, 4d, and 4e; lineage II covering serovars 1/2a, 1/2c, 3a, and 3c; and lineage III containing serovars 4a and 4c.9,10 More recently, it has been shown that L. monocytogenes lineage III can be further classified into three distinct genetic subgroups IIIA, IIIB, and IIIC, with subgroup IIIA being composed of typical rhamnose-utilizing avirulent serovar 4a (which react with inlA and lmo0733, but not with inlJ and inlC, lmo1134, and lmo2672 primers) and virulent serovar 4c strains (which react with inlA, lmo0733, inlJ, and/or inlC, but not with

207

208

Molecular Detection of Foodborne Pathogens

Table 15.1 Compositions of Somatic (O)/Flagellar (H) Antigens in Listeria Serovars

Table 15.2 L. monocytogenes Serovars Causing Human Listeriosis

Serovar

Serotype

1/2a 1/2b 1/2c 3a 3b 3c 4a 4ab 4b 4c 4d 4e 7 5 6a 6b

O-antigen I, II I, II I, II II, IV II, IV II, IV (V), VII, IX V, VI, VII, IX V, VI V, VII (V), VI, VIII V, VI (VIII), (IX) XII, XIII (V), VI, (VIII), X V, (VI), (VII), (IX), XV (V), (VI) (VII), IX, X, XI

H-antigen A, B A, B, C B, D A, B A, B, C B, D A, B, C A, B, C A, B, C A, B, C A, B, C A, B, C A, B, C A, B, C A, B, C A, B, C

Source: Adapted from Seeliger, H.P.R. and Jones, D., Listeria. In Bergey’s Manual of Systematic Bacteriology, vol. 2. pp. 1235–1245. Sneath, P.H.A. et al. (eds.). Williams and Wilkins, Baltimore, MD, 1986.

lmo1134 and lmo2672 primers); subgroup IIIC of atypical nonrhamnose-utilizing virulent serovar 4c strains (which react with inlA, lmo0733, inlJ, inlC, and lmo2672, but not with lmo1134 primers); and subgroup IIIB of atypical rhamnose-negative virulent nonserovar 4a and nonserovar 4c strains (which react with inlA and lmo2672 and inlC, but not with lmo0733, inlJ, and lmo1134 primers), some of which may be related to serovar 7 (which also react with ORF2819 primers).11,12 It is possible that subgroup IIIB (including serovar 7) may represent a novel subspecies within L. monocytogenes.11,12 In accordance with its virulence potential in mammalian hosts, L. monocytogenes is also differentiated into pathogenic and nonpathogenic strains. It is most notable that L. monocytogenes serovars 4b, 1/2a, 1/2b, and 1/2c are responsible for over 98% isolations from clinical cases of human listeriosis, with serova 4b alone accounting for 49% of Listeria-related foodborne diseases (Table 15.2).13 In experimental mouse models, L. monocytogenes serovars 4b, 1/2a, 1/2b, and 1/2c tend to display higher infectivity than other serovars through intragastric inoculation.14 However, all L. monocytogenes serovars except 4a have the capability to cause mouse mortality via intraperitoneal route (Table 15.3).15–17 In general, Listeria spp. are nutritionally undemanding and grow well on a number of nonselective microbiological media such as tryptone soy broth and brain heart infusion (BHI) broth, although the availability of biotin, riboflavin, thiamine, thioctic acid, cysteine, glutamine, isoleucine, leucine, and valine as well as carbohydrates (e.g., glucose) aids to their optimal growth. The colonies (measuring 0.5–1.5 mm in diameter) of Listeria spp. usually emerge after 24–48 h incubation on laboratory agar media (e.g., blood agar or BHI

4b 1/2a 1/2b 1/2c 3a/3b

No. of Isolates (%) 294/603 (49%) 163/603 (27%) 120/603 (20%) 22/603 (4%) 4/603 (<1%)

Tendency to Cause CNS infections > M/N diseases > Bacteraemia Bacteraemia > M/N diseases > CNS infections M/N diseases > Bacteraemia > CNS infections Bacteraemia > CNS infections > M/N diseases Bacteraemia

M/N diseases, maternal-neonatal diseases; CNS infections, central nerve system infections. Source: Adapted from Goulet V., Euro Surveill., 11, 79, 2006, which was based on the analysis of 603 L. monocytogenes isolates from 603 French patients during 2001–2003. Notes:

agar), and appear as round, translucent, low convex with a smooth surface and a crystalline center. After 3–7 days, older colonies become larger (3–5 mm in diameter) and often show a more opaque appearance. Upon extended incubation, some rough colonies with a sunken center may develop occasionally; and some Listeria cells from older cultures may lose their ability to retain the Gram stain, and fail to display characteristic violet color. The optimal growth temperatures for Listeria are between 30 and 37ºC; at 20–28ºC, Listeria are motile with peritrichous flagella, but become nonmotile at 37°C. Nonetheless, due to their psychrotrophic nature, Listeria are able to survive and grow below 0ºC and up to 45°C. In terms of their pH resistance, Listeria grow well between pH 6 and pH 9, and show remarkable ability to recover after treatment with 100 mM Tris buffer at pH 3 and pH 12.5,18 In addition, Listeria can tolerate a range of osmolarity and have been shown to grow in the presence of 10% NaCl and recover from a 20-h treatment with 7 M NaCl.18

15.1.2  Epidemiology Due to their renowned ability to withstand extreme pH, temperature, and salt ranges, Listeria spp. are widely distributed in all environments. Although Listeria spp. are rarely isolated from unpolluted seawater, ground-water and spring (with an average presence of about 1%) where the availaibilty of nutrients is limited, they are commonly found in river and bay waters, sewage, and industrial and farming effluents (with an average detection rate of ≥ 20%). Indeed, the incidence of Listeria spp. in water appears to increase with animal and human activity, as polluted water and sewage/sludge contain various nutrients essential for Listeria growth in compassion with unpolluted water. Listeria spp. are also routinely isolated in soil and plants (with an average presence of 20%) where they exist as saprophytes and sanvengers. L. monocytogenes (especially serovars 1/2a, 1/2b, and 1/2c, which happen to be commonly associated with human foodborne listeriosis) has the ability to adhere to the surfaces of food processing

209

Listeria

Table 15.3 L. monocytogenes Serovars and Relative Virulence PCR Strain

Source

Serovar

inlJ

inlC

LD50a

Relative Virulence (%)b

1

+

+

<7.0 × 10

70

1/2a

+

+

<1.1 × 107

100

2

+

+

1.6 × 109

30

Ruminant brain

4a

–

–

1.9 × 1010

0

HCC25

Catfish kidney

4a

–

–

3.5 × 1010

0

ATCC 19115

Human

4b

+

+

6.0 × 108

70

ATCC 19116

Chicken

4c

+

–

2.6 × 108

100

874

Cow brain

4c

+

+

<8.0 × 107

100

ATCC 19117

Sheep

4d

+

+

8.8 × 108

40

ATCC 19118

Chicken

4e

+

+

7.8 × 109

50

R2-142

Food

7

–

+

<5 × 107

100

HCC8

Catfish brain

EGD

Guinea pig

ATCC 19112

Human

ATCC 19114

8

Sources: Adapted from Liu, D., FEMS Microbiol. Lett., 233, 159, 2004 and Liu, D. et al., J. Clin. Microbiol., 44, 2229, 2006. LD50 (medium lethal dose) values were determined in A/J mouse strain via intraperitoneal route.15,16 b Relative virulence (%) is calculated by dividing the number of dead mice with the total number of mice tested using EGD as reference.16

a

benches, machineries, and floors, in which it develops into biofilm matrix with increased resistance to adverse conditions. On the whole, L. monocytogenes and especially L. innocua are more frequently isolated from the environments than L. seeligeri, L. ivanovii, and L. seeligeri; and L. grayi is seldom reported from environmental samples. Similarly, Listeria spp. are isolated more frequently from fish and seafoods caught in waters with farms and human settlements nearby than those harvested from unpolluted waters. In addition, a varity of meat products (e.g., chicken, pork, and beef) provide nutritional media for Listeria growth and maintenance, and it is no surprise that a relatviley high prevalence (often in the range of 20–65%) of L. monocytogenes and Listeria spp is frequently observed in these products. On the other hand, the presence of Listeria spp. and L. monocytogenes in raw bulk tank milk, dairy milk and vegetables is generally low (often <5%). Considering the tendency toward increased comsuption of processed and ready-to-eat (RTE) food products in recent decades, contaminated foods (e.g., “gravad” or smoked rainbow trout, smoked mussels, RTE meat, and dairy products) have been responsible for many of the human listeriosis outbreaks documented worldwide. This can be attributable to the fact that vacuum packaging, cooling, acid, and alkaline treatments may eliminate other bacteria in the manufactured food products but not Listeria spp., since Listeria are capable of producing fatty acids of the anteiso-type instead of the iso-type to enhance their resistance to cold temperature and other external stresses. While between 15 and 20% of wildlife and domestic animals are found to be infected with Listeria spp. (including pathogenic L. monocytogenes and L. ivanovii) via ingestion of Listeria-contaminated feed (e.g., silage and grasses) and water, the prevalence of Listeria spp. in human population is relatively low at about 3.4–7.7 cases of listeriosis per million as reported in the USA and France. However, the incidence of L. monocytogenes infections is notably higher among

pregnant women (accounting for about 17–24% of clinical cases of listeriosis) and elderly (50% of the cases involving people aged >70 years) as well as individuals with underlying diseases (e.g., cancer, organ transplantation, AIDS, chronic hepatic disorder and diabetes), who are taking medications (e.g., systemic steroids or cyclosporine), or receiving radiation treatment or chemotherapy. The prevalence of Listeria spp. in asymptomatic healthy adults ranges from 1 to 5%. Interestingly, of the 603 human listeriosis cases recorded in France during 2001–2003 in France (see Table 15.2), serovars 4b, 1/2a, and 1/2b accounted for 96% of the cases.13

15.1.3  Clinical Features L. monocytogenes (originally named Bacterium monocytogenes) was first identified in the UK by E.G.D. Murray in 1926 as a cause of infection with monocytosis in laboratory rodents. For a considerable period, this bacterium has been regarded largely as a bacterial pathogen of animals (e.g., ruminants, pigs, dogs, and cats), with the affected animals showing encephalitis signs (with uncoordinated posture and circular movement, the so-called “circling disease”), abortion or septicaemia. Along with a trend toward increasing consumption of RTE and heat-and-eat convenient food products, L. monocytogenes has emerged as a significant foodborne pathogen in the late 1970s and early 1980s, when several large outbreaks of foodrelated human listeriosis cases with significant morbidity and mortality occurred. In fact, with mortality rates on average approaching 30%, L. monocytogenes far exceeds other common foodborne pathogens, such as Salmonella enteritidis (with a mortality of 0.38%), Campylobacter species (0.02–0.1%) and Vibrio species (0.005–0.01%) in terms of disease severity.19,20 During the early stages of infection, human listeriosis often displays nonspecific flu-like symptoms (e.g., fever and chills,

210

fatigue, headache, and muscular and joint pain) and gastroenteritis (which may be observed before the onset of bacteraemia or in healthy individuals infected with a high dose of L. monocytogenes not accompanied by other symptoms). The severity and prognosis of the disease are clearly dependent on the immune status of individuals. In the immunocompetent individuals, the invading L. monocytogenes bacteria are valantly fought off and eliminated by host’s cell-mediated immune network and the disease symptoms often disappear with a short period (in a matter of days). However, in the immunocomprimised individuals such as pregnant women, neonates, the elderly, and individuals under chemotherapy that suppresses immune functions, L. monocytogenes has a propensity to cause especially severe problems with abortion and occasional death as frequent outcomes, given L. monocytogenes’ ability to cross host’s intestinal, blood-brain, and fetoplacental barriers, affecting the uterus at pregnancy, the central nervous system or the bloodstream. In early-onset fetomaternal/neonatal listeriosis, L. monocytogenes enter the blood stream of pregnant women (who may be asymptomatic), colonize the placenta, and invade the fetus, leading to abortion within the last third of the gestation period, with a stillborn fetus or the birth of a baby showing a severe, most often fatal septicemic syndrome known as “granulomatosis infantiseptica.” In the late-onset perinatal listeriosis resulting either from low-level transplacental infection or horizontal transmission at neonatology wards, newborns between weeks one and eight postpartum often show a febrile syndrome associated with meningitis, although gastroenteritis and pneumonia can also be observed. The mortality of later-onset listeriosis is generally lower than that in early-onset listeriosis.

15.1.4  Pathogenesis In a typical mode of infection, L. monocytogenes enters the host via contaminated foods. After sustaining exposure to host proteolytic enzymes, the acidic stomach environment (pH 2.0), bile salts and nonspecific inflammatory attacks, largely through the actions of several stress-response genes (opuCA, lmo1421, and bsh) and related proteins.4 L. monocytogenes adheres to and is internalized by host cells with the assistance of a family of surface proteins called internalins. The most notable internalins are InlA and InlB. Whereas InlA (an 88-kDa protein encoded by inlA) interacts with E-cadherin to mediate L. monocytogenes entry into epithelial cells, InlB (a 65-kDa protein encoded by inlB) recognizes C1q-R (or Met) to facilitate L. monocytogenes entry into a much broader range of host-cell types, including hepatocytes, fibroblasts, and epithelioid cells. Besides InlA and InlB, other internalins (e.g., InlC and InlJ) also appear to be involved in the later (i.e., post intestinal) stages of L. monocytogenes infection.21–23 This internalization process enhances L. monocytogenes evasion of host immune surveillance functions.7 Following its uptake by host cells, L. monocytogenes is primarily located in single-membraned vacuoles. Two virulence-associated molecules are responsible for lysis of the

Molecular Detection of Foodborne Pathogens

primary single-membraned vacuoles and subsequent escape by L. monocytogenes: listeriolysin O (LLO) and phosphatidylinositol-phospholipase C (PI-PLC). LLO (a 529-amino acid, 58-kDa protein encoded by hly) is a pore-forming, thiol-activated toxin that is essential for L. monocytogenes virulence. PI-PLC (a 33-kDa protein encoded by plcA), acting in synergy with phosphatidylcholine-phospholipase C (PC-PLC, a 29-kDa protein encoded by plcB), aids LLO in lysing the primary vacuoles.7 After lysis of the primary single-membraned vacuoles, L. monocytogenes is released to the cytosol, where it starts multiplying intracellularly shortly afterward (with generation time of 40–60 min, which is comparable to those observed in rich broth culture). The host cell cytoplasm hence allows listerial growth with high efficiency. The intracellular mobility and cell-to-cell spread of L. monocytogenes require another surface protein, ActA (a 67-kDa protein encoded by actA), which is cotranscribed with PC-PLC and mediates the formation of polarized actin tails that propel the bacteria toward the cytoplasmic membrane. At the membrane, bacteria become enveloped in filopodium-like structures that are recognized and engulfed by adjacent cells, resulting in the formation of secondary double-membraned vacuoles. A successful lysis of the secondary double-membraned vacuoles signals the beginning of a new infection cycle, which is dependent on PC-PLC upon activation by Mpl (a 60-kDa metalloprotease encoded by mpl).7 The genes encoding the virulence-associated proteins PI-PLC, LLO, Mpl, ActA, and PC-PLC are located in a 9.6-kb virulence gene cluster,23 which is primarily regulated by a pleiotropic virulence regulator, PrfA, (a 27-kDa protein encoded by prfA). The prfA gene is situated immediately downstream of, and sometimes cotranscribed with, the plcA gene. PrfA activates the transcription of many L. monocytogenes virulence-associated genes. The genes encoding InlA and InlB are positioned elsewhere in the genome. As the inlA and inlB genes possess a transcription binding site similar to that recognized by PrfA, they may also be partially regulated by PrfA. In addition to these virulence-associated genes and proteins, many other genes such as iap (encoding invasion-associated protein, or Iap, now known as p60) also play essential roles in L. monocytogenes virulence and pathogenicity.7 Belonging to a protein family with two other members in L. monocytogenes (p45, a peptidoglycan (PG) lytic protein encoded by the spl gene, and the hypothetical protein encoded by lmo394), p60 is a highly basic protein of 484 amino acids with an indirect role in modifying bacterial behavior via its impact on cell division.

15.1.5  Genomics The recent completion of the whole genome sequences of several Listeria strains has uncovered valuable new details on the differential genetic compositions among Listeria species (Table 15.4). Currently, the complete genome sequences of two L. monocytogenes serovar 1/2a (EGD-e and F6854) and two serovar 4b (F2365 and H7858) strains, one L.

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Table 15.4 Comparison of Listeria Genomes L. monocytogenes Feature Serovar Genome size (bp) G+C content (%) Protein-coding genes (ORFs) Strain-specific genes

EGD-e

F6854

F2365

H7858

L. innocua CLIP 11262

L. welshimeri SLCC 5334

1/2a 2,944,528 39 2853 61

1/2a 2,950,285 37.8 2973 97

4b 2,905,187 38 2821 51

4b 2,972,254 38 3024 69

6a 3,011,208 37.4 2973 149

6b 2,814,130 36.4 2780 311

Sources: Adapted from Glaser, P. et al., Science, 294, 849, 2001; Nelson, K.E. et al., Nucleic Acids Res., 32, 2386, 2004 and Hain, T. et al., J. Bacteriol., 188, 7405, 2007.

innocua strain (CLIP 11262) and one L. welshimeri strain (SLCC 5334) are available (Table 15.4).1–3 The sequencing analyses of 15 other L. monocytogenes strains covering serovars 1/2a (7), 1/2b (4), 1/2c (1), 4b (1), and unknown (2) are in progress. Comparison of the six Listeria genomes at the nucleotide and predicted protein levels shows that in addition to many shared genetic components, a total of 51, 97, 69, and 61 strain-specific genes are identifiable from L. monocytogenes serovar 1/2a strains EGD-e and F6854, and serovar 4b strains F2365 and H7858, respectively (Table 15.4). Further analysis reveals that 83 of these genes are limited to the serovar 1/2a strains (of lineage II), and 51 genes are limited to the serovar 4b strains (of lineage I). In addition, 149 and 311 speciesspecific genes are recognized in L. innocua CLIP 11262 and L. welshimeri SLCC 5334, respectively (Table 15.4).1–3 Since L. monocytogenes strain-specific genes tend to have atypical base composition, some of them may have been acquired by horizontal gene transfer. Moreover, some L. monocytogenes strain-specific genes encode putative surfaceassociated proteins such as internalins, which may contribute to the increased pathogenicity of the strains concerned.2 Surprisingly, 37 (44%) of the 83 serovar 1/2a-specific and 33 (65%) of the 51 serovar 4b-specific genes encode hypothetical proteins for which no biochemical information is available. The serovar 1/2a-specific genes include three clusters that encode pathways for the transport and metabolism of carbohydrates, an operon that encodes the biosynthetic pathway for the antigenic rhamnose substituents that decorate the cell wall-associated teichoic acid polymer in serovar 1/2a strains, five glycosyl transferases and an adenine-specific DNA methyltransferase.2 While the genomes of serovar 4b strains (F2365 and H7858) do not contain intact insertion sequence (IS) elements, they harbor four copies of transposase ORFA of the IS3 family. Similarly, the genomes of serovar 1/2a strains (F6854 and EGD-e) possess three copies of the same transposase ORFA in a corresponding location. The extra copy of the transposase ORFA in the serotype 4b strains F2365 and H7858 may have resulted from a duplication event (along with the associated regions). Further, an intact IS element (ISLmo1) is present in the serotype 1/2a strains F6854 (two

copies) and EGD-e (three copies), respectively. Because all L. monocytogenes genomes sequenced to date harbor copies of transposase ORFA, it is probable that transposase ORFA gene may have originated from the genome of an ancestral Listeria before the strains diverged, in comparison to ISLmo1, which may represent a recent acquisition in serovar 1/2a strains.2 Apart from possessing many unique genes, Listeria species/serovars also demonstrate differences among their shared genes. For example, Southern blot analysis with a virulencespecific internalin inlJ probe revealed that genomic DNA from the more pathogenic L. monocytogenes serovars (e.g., 1/2a, 1/2b, 1/2c, and 4b) tends to show a 5.0-kb HindIII band, and that from the less pathogenic serovars (e.g., 3a and 4c) forms a 1.5- or 2.0-kb HindIII fragment. On the other hand, genomic DNA from the nonpathogenic L. monocytogenes serovars (i.e., 4a) shows no band.12,24 Similarly, although being present in all L. monocytogenes serovars, transcriptional regulator gene lmo0733 is recognized as a HindIII band of 5.0 or 6.0 kb in the more pathogenic serovars (e.g., 1/2a, 1/2c, and 4b) and as a HindIII band of 1.0 or 1.5 kb in the less pathogenic or nonpathogenic serovars (e.g., 4a and 4c).12,24 In addition, examination of PG N-deacetylase gene pgdA (lmo0415) in Listeria by Southern blot indicates that L. monocytogenes serotypes 4a and 4c show a smaller band of 1.5 kb while other serotypes have a larger band of 4.0 or 5.0 kb. Further, whereas L. innocua and L. welshimeri display a band of 4.0 kb, L. seeligeri a band of 3.0 kb, L. ivanovii a band of 1.5 kb, L. grayi forms no band at all.12 Given that N-deacetylation of PG provides strength and rigidity to bacterial cell wall, it is no surpise that the absence of pgdA (lmo0415) in L. grayi renders it susceptible to lysozyme treament.12 Indeed, a L. monocytogenes mutant lacking PG N-deacetylase gene (i.e., pgdA or lmo0415) is also highly sensitive to killing by lysozyme, and has impaired ability to survive and/or replicate within macrophages.25 In addition, recent studies indicate that there are some unusual Listeria strains possessing genetic elements that normally belong to other Listeria species. For example, examination of several hemolytic L. innocua strains (e.g., PRL/NW 15B95 and J1-023) revealed the presence of Listeria pathogenicity island 1 (LIPI-1, or PrfA-regulated virulence gene

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cluster), in these atypical L. innocua strains, contributing to their positive hemolytic reactions.26,27 The PrfA-regulated virulence gene cluster was previously found in L. monocytogenes, L. ivanovii, and L. seeligeri only.23 Futhermore, these atypical L. innocua strains also harbor L. monocytogenes internalin gene inlA and partial internalin gene inlB sequences.26,27 The L. innocua species status of these strains was only resolved after DNA–DNA hybridization, microarray, and PCR experiments targeting a host of L. innocua-specific genes.26,27 Similarly, L. monocyotogenes serovars 4a and 4c strains harboring L. innocua-specific genes lin0464, lin0372, and lin1073 have also been described.28–30 These findings suggest that L. innocua may have descended from L. monocytogenes through successive gene deletion, and that atypical hemolytic strains PRL/NW 15B95 and J1-023 may represent some intermediates that have yet to completely eliminate or lose the remaining L. monocytogenes genes.27 Doumith et al.31 also presented evidence that an ancestral L. monocytogenes strain (probably of serovar 1/2a) might have evolved to become serovar 4b through gene deletion, which latter gave rise to serovar 4a possibly via serovar 4c, and finally to L. innocua via additional gene deletion events. The fact that L. monocytogenes serovar 4c and 4a strains harboring L. innocua-specific genes lin0464, lin0372, and lin1073 indicates the possibility that during their transition to L. innocua, some L. monocytogenes strains may pick up certain L. innocua genes while shedding their own. Obviously, additional microarray and sequencing studies are required to determine what other L. innocua genes that have been gained by these unusual L. monocytogenes 4a and 4c strains, and what L. monocytogenes genes have been lost by these strains. The availability of these data will lead to a better understanding of Listeria evolution and yield new details on the phylogenetic relationship among individual Listeria species.

15.1.6  Diagnosis 15.1.6.1  Conventian Techniques (i) In vitro culture isolation: As Listeria are not distinguishable from other Gram-positive bacteria, and are often present in insufficient numbers for direct detection by serological and molecular procedures, in vitro culture enrichment and isolation are undertaken. For isolation of Listeria spp. from nonsterile samples such as food, environmental samples and feces, a range of selective broths and agars are available. PALCAM and OXFORD selective agars are recommended by ISO 11290-1, 2, for primary and secondary plating media, respectivley, for isolation of all Listeria spp.32 Furthermore, chromogenic medium ALOA is also useful for Listeria isolation. This agar medium incorporates 5-bromo-4-chloro3-indolyl-β-D-glucopyranosid as an enzyme substrate for β-D-glucosidase that is secreted by all Listeria spp, leading to the formation of typical blue-turquoise colonies. The inclusion in ALOA of L-α-phosphatidylinositol as a second substrate for phospholipase C, which is a virulence factor produced by pathogenic L. monocytogenes and L. ivanovii, results in the formation of typical blue-turquoise colonies with

Molecular Detection of Foodborne Pathogens

an outer white zone of precipitation. Application of selective chromogenic L. monocytogenes plating media facilitates the presumptive detection of L. monocytogenes within 24 or 48 h. Subsequent confirmation of Listeria identity can be then achieved with biochemical, immunological, and molecular tests. (ii) Biochemical tests: Besides being catalase positive, and indole and oxidase negative, Listeria species can hydrolyse aesculin, but not urea. Additionally, Listeria species show significant variations in their ability to hemolyse horse or sheep red blood cells, and in their ability to produce acid from L-rhamnose, D-xylose and α-methyl-D-mannoside. Of the six Listeria species, L. monocytogenes (producing listeriolysin or LLO), L. ivanovii (producing ivanolysin or ILO), and L. seeligeri (producing seeligeriolysin or SLO) are known to be hemolytic, while L. innocua, L. welshimeri, and L. grayi are generally not. This difference in hemolytic activity is utilized as one of the criteria for discrimination among Listeria species. Furthermore, L. ivanovii is differentiated biochemically from L. monocytogenes and other Listeria species by its production of a wide, clear or double zone of haemolysis on sheep or horse blood agar, a positive Christie–Atkins–Munch-Petersen (CAMP) reaction with Rhodococcus equi but not with hemolytic Staphylococcus aureus, and fermentation of D-xylose but not L-rhamnose.33 Additionally, L. ivanovii is distinguished from L. monocytogenes by its strong lecithinase reaction with or without charcoal in the medium, in comparison to L. monocytogenes, which requires charcoal for its lecithinase reaction.34 Similarly, L. innocua is distinguished from L. monocytogenes on the basis of its negative CAMP reaction and its failure to cause β-haemolysis or to show PI-PLC activity on chromogenic media. L. welshimeri is distinguished from other Listeria species by its negative β-haemolysis and CAMP reactions, and by its acid production from D-xylose and α-methyl-Dmannoside.35 The development and application of several commercial kits (e.g., API-Listeria, Listeria ID, Microbact 12L, and MICRO-ID Listeria) further facilitate the concurrent and convenient biochemical identification of L. monocytogenes and differentiation from other Listeria spp. (iii) Serological tests: Agglutination and enzyme-based immunoassay (EIA) employing polyclonal and monoclonal antibodies to Listeria antigens are examples of serological methods that provide an ease-of-use approach for speciesspecific identification of L. monocytogenes and other Listeria species. A large number of commercial kits on the basis of serological reactivity have been marketed by various companies. These include Assurance Listeria EIA, Dynabeads antiListeria, EIA Foss Listeria, ListerTest Listeria Rapid Test, Listeria-Tek, TECRA Listeria Visual Immuno Assay, Transia Plate Listeria, Vidas LIS, Vidas LMO, and VIP for Listeria, etc. In additon to species-specific identification, serological techniques are also useful for serotyping Listeria strains.36 Specifically, the presence of unique combinations of somatic (O) antigens of I–XV and flagellar (H) antigens of A–D in Listeria strains enables their classification into 15 serovars (Table 15.1).8 This information is critical for epidemiological

213

Listeria

tracking of L. monocytogenes involved in epidemic outbreaks of listeriosis. However, with both L. monocytogenes and L. seeligeri containing serovars 1/2a, 1/2b, 3b, 4a, 4b, 4c, and 6b, the inability of serotyping methods to correlate serovars directly with species identities limits their potential for clinical application. This is further exacerbated by the fact that due to antigen sharing among various L. monocytogenes serovars, with 1/2a and 3c both possessing H antigens A and B; 4a–d, 1/2b and 3b all having H antigens A, B, and C; 1/2c and 3a both harboring H antigens B and D; and multiple, common O antigens being present in different serotypes (Table 15.1), it can be a challenge to conclusively determine the serovar of some L. monocytogenes strains, especially among serovars 4a, 4b, and 4c, which demonstrate contrasting pathogenicity.24 The high cost of acquiring subtype-specific antisera also makes the use of serotyping methods unattractive. Like biochemical methods, serotyping methods are also liable to give occasional discrepant results because of their dependence on the phenotypic characteristics of Listeria bacteria. For these reasons, serotyping methods have largely been superseded by molecular procedures that are intrinsically more specific and sensitive for the identification and differentiation of Listeria species. (iv) In vivo assays: Because L. monocytogenes serovars demonstrate varied infectivity, pathogenicity, and virulence in mammalian hosts, in vivo assays based on animal models such as rodents offer a valuable means to confirm their virulence potential as well as their identity in light of the fact that L. innocua, L. seeligeri, L. welshimeri, and L. grayi are nonpathogenic. Interestingly, through intragastric injection, L. monocytogenes epidemic strains appeared to show higher invasiveness than environmental strains,37 although via intraperitoneal inoculation, all L. monocytogenes serovars but 4a were capable of causing mouse mortality (see Table 15.3).15– 17,38,39 Moreover, in vivo mouse models provide an indispensable tool for the studies of L. monocytogenes pathogenesis and virlence mechanisms that are not easily obtained with in vitro techniques. 15.1.6.2  Molecular Techniques While conventional techniques have proven useful for identification of Listeria bacteria, they often suffer from the shortcomings such as time-consuming and variability due to their dependence on the phenotypic characteristics of the bacteria. On the other hand, molecular procedures are intrinsically more specific and sensitive, and have been increasingly regarded as the methods of choice for improved identification and differentiation of Listeria species. A number of molecular characteristics of Listeria bacteria have been exploited for their identification. For example, the proportion of guanine and cytosine (or G+C content, usually expressed as a mol%) among Listeria species differs from each other (ranging from 36 to 38 mol%), and also from other bacteria (e.g., Esherichia coli at about 50 mol%). Random primer sites and repetitive elements (e.g., repetitive extragenic palindromes or REP; enterobacterial repetitive intergenic consensus sequences or ERIC; variable-number tandem repeats or

VNTR) offer useful targets for PCR-based amplification and differentiation, which are not depedent on the availability of detailed gene sequences of interest. Additionally, restriction enzyme sites provide a way to differentiate and type Listeria strains using pulse-field gel electrophoreis (PFGE), restriction fragment length polymorphism (RFLP) and ribotyping procedures, etc.40 More specific gene targets for Listeria identification comprise rRNA coding genes (i.e., small-subunit 16S and large subunit 23S rRNA genes as well as region 16S–23S intergenic spacer), house keeping genes (groESL, rpoB, recA, gyrB, and prs), genes encoding for invasion associated protein (iap), flagellin A (flaA) and fibronectin-binding protein (fbp), a large number of species-specific (e.g., hly, plcA, plcB, actA, mpl, inlA, inlB, delayed-type hypersensitivity gene, aminopeptide gene and putative transcriptional regulator gene lmo0733), and group-specific genes (e.g, lmo2821, lmo1134, and lmo2672).15,40,41 In addition, a number of genes unique to individual Listeria species have been also described and applied for improved differentiation of nonmoncytogenes Listeria species from L. monocytogenes.29,42–84 Incorporation of specific primers from some of these genes facilitated the development of multiplex PCR assays for improved differentiation among Listeria species.28 Further, specific primers have also been applied in multiplex PCR formats for separation of L. monocytogenes serovar groups.85,86 In addition, by using primers from inlA for species-specific recognition, and those from inlJ (or lmo2821) and inlC for virulence determination in a multiplex PCR, it is possible to differentiate L. monocytogenes naturally avirulent serovar 4a strains from other serovars with potential to cause mouse mortility via intraperitoneal route.17,87,88 The advent of microarray technology in recent years provides new opportunities for L. monocytogenes gene activation analysis in host-pathogen relationships, without the limitations that are encountered with the existing genomic and transcriptional procedures. In other words, the microarray technology combines the feature of genomic technique for confirming the presence of genes with that of transcription analysis for evaluating gene expression. This permits the investigation of the in/activation (down/up-regulation) of L. monocytogenes virulence-associated genes during distinct phases of infection in mammalian hosts and under other strenuous external conditions.89,90

15.2  Methods 15.2.1  Sample Preparation Bacterial isolation. As most food samples contain very few L. monocytogenes bacteria, an enrichment, and/or isolation process is required prior to identification by biochemical and molecular techniques. Typically, 25 g of food is homogenized using a blender in 225 ml of culture medium (e.g., half Fraser broth, which contains half a volume of acriflavine and nalidixic acid), and incubated at 30°C for 24 h. Afterward, a 0.1 ml of the primary enriched culture is added

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Molecular Detection of Foodborne Pathogens

to 10 ml of full strength Fraser broth (which contains one volume of acriflavine and nalidixic acid), and incubated at 37°C for 24 h. A loopful of the secondary enrichment culture is then streaked onto BCM® Listeria monocytogenes plating medium at 37°C for 24 h for production of turguoise colonies.32

Alternatively, if Listeria DNA is intended for PCR only, it can be prepared by boiling. Namely, suspend one to two colonies in 100 μl of 50 mM NaOH, vortex, heat at 100°C for 10 min, add 10 µl of 1 M Tris pH 8.0, centrifuge at 13,000×g for 1 min, and use 2–5 μl in 25 µl PCR mixture.

DNA extraction:

15.2.2  Detection Procedures

(1) Transfer Listeria colonies (or retrieve isolates from frozen glycerol stocks) onto 5% sheep blood agar plate (e.g., Tryptic soy agar II, Becton Dickinson Microbiology Systems, Cockeysville, MD) and incubate at 37°C overnight. Then transfer several colonies of each strain into 5 ml of BHI broth (Difco Laboratories, Detroit, MI) and keep at 37°C for 16–18 h, shaking at 250 rpm (with OD at 620 nm of >1.0). (2) Pellet the bacteria at 3000 rpm for 10 min, and decant the supernatant. (3) Resuspend the cell pellet in 500 μl of DNA lysis buffer (30 mM Tris-HCl pH 8.0, 30 mM EDTA pH 8.0, 150 mM NaCl, 0.5% SDS, 0.5% Triton X-100 and 0.5% Tween 20) by vortexing; add 10 μl of freshly made lysozyme (10 mg/ml), and 5 μl of RNase A (10 mg/ml), and incubate at 37°C for 30 min. Note: a COmbination of detergents is included in this DNA lysis buffer to allow efficient lysis of all bacterial categories. While most Gram-negative and Gram-positive bacteria are readily lysed in the presence of 1% SDS, some Gram-positive bacteria such as Listeria tend to lyse more easily in the presence of 0.5–1% Triton X-100. (4) Add 10 μl of proteinase K (10 mg/ml), and incubate at 56°C for 1–2 h (when the lysate becomes clear). (5) Transfer the lysate to a 1.5 ml tube, add one volume (500 μl) of phenol/chloroform/isoamyl-alcohol (25:24:1), and mix by inversion. (6) Centrifuge at 13,000 rpm for 10 min (repeat Steps 6 and 7 once for increased DNA purity if desired). (7) Transfer supernatant to a new 1.5-ml tube, add one volume (about 500 μl) of isopropyl alcohol and 20 μl of 5 M NaCl (to a final concentration of 0.10–0.15 M), mix by inversion, and centrifuge at 13,000 rpm for 10 min at 4°C. (8) Discard supernatant, add 300 μl of 70% ethanol (stored at –20°C), and centrifuge at 13,000 rpm for 2 min at 4°C. (9) Discard supernatant, invert the tube on a paper towel, air dry for 10 min. (10) Dissolve the DNA pellet in 200 μl of 1×TE (10 mM Tris-HCl pH 8.0 and 1 mM EDTA pH 8.0), determine the DNA concentration at UV 260/280 nm in a spectrophotometer, and store at –20°C. A small amount of DNA from each strain is diluted in distilled water to 10 ng/µl for PCR analysis.

15.2.2.1 Multipex PCR Identification of Listeria Species Principle: Based on the six Listeria species-specific genes described previously,42–47 Huang et al.28 designed two multiplex PCR, with one focusing on L. welshimeri, L. monocytogenes, and L. seeligeri; and another on L. ivanovii, L. grayi, and L. innocua (Table 15.5) Procedure: (1) Prepare the PCR mixture (25 μl) containing 0.5 U Taq Gold DNA polymerase (Applied Biosystems, Foster City, CA), 1 × PCR buffer (10 mM Tris pH 8.3, 50 mM KCl, 2.5 mM MgCl2), 200 μM dNTPs, 25 pmol each primer (L. welshimeri, L. monocytogenes, and L. seeligeri in pool 1; L. ivanovii, L. grayi, and L. innocua in pool 2), 2 μl DNA (prepared by boiling method), and distilled water (to 25 μl). Include a tube containing PCR mixture without DNA template as a negative control in each run. (2) Run the following cycling program in a GeneAmp PCR System 9700 (Perkin Elmer, Boston, MA): one cycle of 95ºC for 9 min; 30 cycles of: 94ºC for 30 sec, 60ºC for 30 sec, and 72ºC for 1 min; and one cycle of 72ºC for 7 min. (3) After completion of PCR cycles, add 3 μl of 6 × DNA loading buffer into each tube; load 15 μl of PCR product on 2.0% agarose gel containing 0.5 μg/ml ethidium bromide. Include a DNA molecular marker in a spare lane. Run at 5 V per cm gel for about 30 min. Visualize under UV light box and and photograph. Note: in pool 1, L. welshimeri forms a band of 608 bp, L. monocytogenes a band of 453 bp, and L. seeligeri a band of 375 bp; in pool 2, L. ivanovii shows a band of 493 bp, L. grayi a band of 420 bp and L. innocua a band of 250 bp. Upon examination of a collection of 456 Listeria isolates collected from routine food quality monitoring schemes and 62 L. monocytogenes isolates from patients, the multiplex PCR had 100% agreement with biochemical methods.44 Nevertheless, it has been demonstrated recently that the lmo0733 gene encoding transcriptional regulator appears to be absent in L. monocytogenes lineage subgroup IIIB strains, which belong to atypical rhamnose-negative virulent non-4a and non-4c strains, some of which may be related to serovar 7,12 and which have been designated as novel evolutionary lineage IV in a more recent study (Ward et al., unpblished data). Therefore, if it is desirable to detect

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Table 15.5 Identities and Sequences of Listeria Primers for Species-Specific Identification Target Gene

Specificity

lwe7-571 (encoding phosphotransferase system enzyme IIBC) Lmo0733 (encoding transcriptional regulator) lse24-315 (encoding internalin)

L. welshimeri

liv22-228 (encoding N-acetylmuramidase)

L. ivanovii

lgr20-246 (encoding oxidoreductase)

L. grayi

iap (encoding invasion-associated protein)

L. innocua

All L. monocytogenes linages but lineage subgroup IIIB L. seeligeri

Primer Sequences (5′→3′)* Pool 1 TCCCACCATTGGTGCTACTCA TTGGCGTACCAAAGAAATACG CGCAAGAAGAAATTGCCATC TCCGCGTTAGAAAAATTCCA CGCGACGGCTAAAGTACTAA ATTGCTCGCTTTGAAGTCGT Pool 2 (N)15CGAATTCCTTATTCACTTGAGC (N)15GGTGCTGCGAACTTAACTCA CTGCACGATCAAGGTCAATC CGTATTGCGCACCAGTGATA TTGCTACTGAAGAAAAAGCA TCTGTTTTTGCTTCTGTAGC

PCR Product (bp) 608 453 375

493 420 250

*N, random nucleotide.

all L. monocytogenes serovars and strains, primers targeting inlA gene may be employed (see Section 15.2.2.3). 15.2.2.2 Multiplex PCR Differentiation of L. monocytogenes Serogroups Principle: Doumith et al.86 developed a multiplex PCR that incorporates L. monocytogenes lmo0737 gene primers for recongition of serovars 1/2a, 1/2c, 3a, and 3c; lmo1118 gene primers for detection of serovars 1/2c and 3c; ORF2819 primers for serovars 1/2b, 3b, 4b, 4d, and 4e; ORF2110 primers for serovars 4b, 4d, and 4e; and prs primers as an internal amplification control covering all L. monocytogenes serovars (Table 15.6). Procedure: (1) Prepare PCR mixture (100 μl) containing 2 U of Taq DNA polymerase (Roche, Boehringer), 0.2 mM dNTPs, 1 × PCR buffer (50 mM Tris-HCl, 10 mM KCl, 50 mM (NH4)2SO4, 2 mM MgCl2, pH 8.3), and the five primer sets (1 μM each for lmo0737, ORF2819, and ORF2110; 1.5 μM each for lmo1118; and 0.2 μM each for prs (Table 15.5). (2) Perform PCR amplification in a thermocycler for one cycle of 94°C for 3 min; 35 cycles of 94°C for 25 sec, 53°C for 1 min 10 sec, and 72°C for 1 min and 10 sec; and one cycle of 72°C for 7 min. (3) After completion, mix 5 μl of PCR mixture with 3 μl of loading buffer and separate on a 2% agarose gel in a TBE buffer (90 mM Trizma base, 90 mM boric acid, 2 mM EDTA, pH 8.3). (4) Stain the gel with ethidium bromide, and visualize with UV transilluminator. Note: all Listeria species (including serovars 4a and 4c) form a 370-bp fragment with the prs gene primers; serovars 1/2a and 3a show a 691 bp fragment with the lmo0737

primers only; serovars 1/2c and 3c display a 691 bp fragment with the lmo0737 primers and a 906 bp fragment with the lmo1118 primers; serovars 1/2b, 3b, and 7 show a 471 bp fragment with the ORF2819 primers; serovars 4b, 4d, and 4e form a 471 bp fragment with the ORF2819 primers and a 597 bp fragment with the ORF2110 primers.86 15.2.2.3 Multipex PCR Determination of L. monocytogenes Virulence Principle: A multiplex PCR was reported recently that incorporates inlA gene primers for species-specific confirmation (with an 800-bp product), and inlC and inlJ gene primers for virulence determination of L. monocytogenes (with 517 bp and 238 bp bands, respectively) (Table 15.7).17 The inclusion of both the inlC and inlJ gene primers in the multiplex PCR has added advantage in that it provides a double verification of L. monocytogenes virulence for in most cases. The usefulness of this multiplex PCR for virulence determination is supported by the observations that L. monocytogenes virulent strains with the potential to cause mouse mortalities via intraperitoneal inoculation of mice are positive with the inlC and/or inlJ gene primers, while naturally avirulent strains unable to produce mouse mortality are negative with these primers.17 Procedure: (1) Prepare the PCR mixture (25 μl) containing 0.8 U Taq DNA polymerase (Fisher Scientific, Houston, TX), 1 × PCR buffer, 200 μM dNTPs, 40 pmol each of inlA primers, 30 pmol each of inlC primers, 20 pmol each of inlJ primers, 10 ng (1 μl) of DNA, and distilled water (to 25μl). Include a tube containing PCR mixture without DNA template as a negative control in each run. (2) Run the following cycling program in a GeneAmp PCR System 9700 (Perkin Elmer, Boston, MA) or

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Table 15.6 Identities and Sequences of L. monocytogenes Primers for Serogrouping Target Gene

Specificity

lmo0737 (encoding protein ofunknown function) lmo1118 ( encoding protein ofunknown function) ORF2819 (encoding transcriptional regulator) ORF2110 (encoding secreted protein) prs (encoding phosphoribosyl pyrophos- phate synthetase)

L. monocytogenes serovars 1/2a, 1/2c, 3a, and 3c L. monocytogenes serovars 1/2c and 3c L. monocytogenes serovars 1/2b, 3b, 4b, 4d, and 4e L. monocytogenes serovars 4b, 4d, and 4e All Listeria species

Primer Sequences (5′→3′) AGGGCTTCAAGGACTTACCC ACGATTTCTGCTTGCCATTC AGGGGTCTTAAATCCTGGAA CGGCTTGTTCGGCATACTTA AGCAAAATGCCAAAACTCGT CATCACTAAAGCCTCCCATTG AGTGGACAATTGATTGGTGAA CATCCATCCCTTACTTTGGAC GCTGAAGAGATTGCGAAAGAAG CAAAGAAACCTTGGATTTGCGG

PCR Product (bp) 691 906 471 597 370

Table 15.7 Identities and Sequences of L. monocytogenes Internalin Gene Primers for Species- and Virulence Determination Target Gene

Specificity

inlA

All L. moocytogenes serovars

inlC

All L. moocytogenes serovars but 4a and some 4c All L. moocytogenes serovars but 4a and some IIIB

inlJ

Primer Sequences (5′→3′) ACGAGTAACGGGACAAATGC CCCGACAGTGGTGCTAGATT AATTCCCACAGGACACAACC CGGGAATGCAATTTTTCACTA TGTAACCCCGCTTACACAGTT AGCGGCTTGGCAGTCTAATA

other cycler: one cycle of 94ºC for 2 min; 30 cycles of: 94ºC for 20 sec, 55ºC for 20 sec, and 72ºC for 50 sec; and eon cycle of 72ºC for 2 min. (3) After completion of PCR cycles, add 3 μl of 6 × DNA loading buffer into each tube; separate the PCR products on 1.5% agarose gel containing 0.5 μg/ml ethidium bromide. Include a DNA molecular marker in a spare lane. (4) Visualize under UV light box and and photograph using a ChemiImager 5500 documentation system (BSI, Stafford, TX). Note: the species identity of the 29 L. monocytogenes strains was validated through the formation of an 800-bp band with the inlA gene primers, and the virulence potential of these strains was confirmed by the production of 517 bp and/or 238 bp bands with the inlC and inlJ gene primers in the multiplex PCR. Even though L. ivanovii strains also harbor a gene with homology to L. monoctyogenes inlC, and cross-react with L. monoctyogenes inlC gene primers, they can readily excluded by their negative reaction with L. monoctyogenes inlA gene primers included in the multiplex PCR.17 Although the presence of inlC and/or inlJ genes in a given L. monocytogenes strain implies its potential virulence and its ability to cause mouse mortality via the intraperitoneal route, it does not indicate the certainty of the strains harboring these genes to produce disease in humans via the

Nucleotide Positions 94612–94631 95411–95392 107306–107325 107822–107802 188989–189009 189226–189207

PCR Product (bp) 800 517 238

conventional oral ingestion. Considering that only three serotypes (1/2a, 1/2b, and 4b) are isolated from human listeriosis cases, many serotypes (e.g., 1/2c, 3a, 3b, 3c, 4c, 4d, and 4e) that are recognized as having virulence potential by the multiplex PCR may encounter difficulty crossing the host’s intestinal or other barriers during infection. Obviously, for a L. monocytogenes strain to be fully infective to humans via oral ingestion, it requires involvement of many other known and uncharacterized virulence genes and proteins. These undefined virulence genes and proteins require further characterization and examination. Nonetheless, an implication from the multiplex PCR results is that while L. monocytogenes serotypes other than 1/2a, 1/2b, and 4b are not commonly associated with human listeriosis, due presumably to their inability to efficiently cross human intestinal and other barriers, they (with the possible exclusion of serotype 4a) may have the potential to produce disease in humans whose immune functions are compromised, or who are exposed to these serotypes via intravenous transfusions or open wounds. The inlA primers described above can be utilized independently for species-specific confirmation of L. monocytogenes if it is unnecessary to determine the virulence potential with the inJ and inC primers. While the inlA primers recognize all L. monocytogenes serovars and lineages, the lmo0733 primers described in Section 15.2.2.1 do not cover the unusual lineage IIIB strains, which may include serovar 7. It appears that the inlA gene is also more conserved across L. monocytogenes

217

Listeria

serovars than inlB gene, as serovars 4a–4e were not detected with a pair of inlB primers designed from serovar 1/2a strain EGD-e.17 Although some L. monocytogenes strains isolated from nonsympomatic human carriers have been shown to produce truncated InlA protein occasionally,91 the incidence of inlA gene mutation in food-derived isolates is remarkably low.92

15.3 Conclusions and Future Perspectives Given its widespread occurrence and its capacity to withstand extreme external stresses such as extreme temperature, pH, and salt ranges, which often underscore many food processing routines, L. monocytogenes has become an increasingly important foodborne pathogen in recent decades. Considering the fact that the early clinical manifestations of human listeriosis are generally nonspecific, and that it can evolve quickly into life-threatening illness in vulnerable population groups (e.g., infants, pregnant women, elderly, and immunocompromised persons) without prompt antibiotic therapy, it is critical to develop a capability to rapidly, sensitively, and reliably detect, identify, and differentiate the pathogen concerned, and to promptly initiate appropriate control and prevention measures against the disease. Although conventional, phenotype-based tests are relatively slow and variable at times, they have played essential roles in the laboratory idenfication and diagnosis of L. monocytogenes and in the tracking of endemic and sporadic listeriosis outbreaks. Following the advent of new generation molecular methods, especially in vitro template amplification techniques such as PCR, speedy, and precise identification of L. monocytogenes has become a reality. While a large number of PCR-based assays targeting various genes have beem reported and proven valuable for discrimination of L. monocytogenes from other closely-related bacteria,42–84 the development and application of multiplex PCR has made species-, serogroupand virulence-specific determination of this bacterium possible and in a much more efficient manner.17,28,85,86 Furthermore, the increasing adoption of real-time PCR platform in the Listeria diagnosis provides unprecedented level of testing convenience and result availability. This also offers the opportunity to quantitate the organisms in the clinical, food, and environmental specimens, and contribute to the improved monitoring of the efficiency of chemotherapy and quality control measures in the food manufacturing facilities and environmental surveillance. In contrast to an increasing sophistication in the nucleic acid amplification and detection technologies, the progress in the development of sample preparation procedures prior to enzymatic amplification has been relatively slow.93 The fact that food samples come in different variety and complexicity, and many components in these samples (e.g., phenolics, glycogen, calcium ions, fat, and other organic substances) are inhibitory to DNA polymerase may have contributed to the inability of developing and optimizing rapid, efficient, and multi-purpose sample processing procedures applicable to all types of food samples. In addition, because many food

samples contain relatively few organisms of interest, which may be below the detection limit of most current molecular techniques, there is need to enrich and concentrate the target organisms before testing. While many concentration procedures based on filtration and centrifugation protocols have been designed and described for individual specimen types, there has been a lack of standardized and optimized procedures for dealing with the myriad type of food specimens. Similarly, although several selective enrichment media (i.e., ISO 11290, FDA BAM and USDA) incorporating acriflavine, nalidixic acid and cycloheximide are in common use, their application seems to be limited to the types of food samples examined, with the USDA method being suited for meat products and the FDA method for other food products (e.g., dairy products, seafood, and vegetables). Clearly, further research in the prior sample handling and processing is to maximize the performance of molecular assays for detection and identification of L. monocytogenes bacteria from various types of samples. This may include the determination of the optimal types and strengths of enrichment media or length of enrichment for individual specimen types containing low number of organisms of interest, and investigation of the most appropriate combination of physical separation and concentration procedures for various types of specimens prior to the enrichment step. The availability of rapid, efficient, and reliable sample preparation procedures is crucial for further strengthening the performance of the molecular tests for direct detection and identification of L. monocytogenes from clinical, food, and environmental specimens. This will not only help shorten the time from specimen submission to result finalization, but also benefit the control and prevention program against listeriosis.

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16 Micrococcus

Friederike Hilbert and Hans-Jürgen Busse University of Veterinary Medicine Vienna

Contents 16.1 Introduction.................................................................................................................................................................... 221 16.1.1 Classification of the Genus Micrococcus.......................................................................................................... 221 16.1.2 Biology, Pathogenesis, and Medical Importance.............................................................................................. 222 16.1.3 Identification and Diagnosis of the Genus Micrococcus.................................................................................. 223 16.2 Methods.......................................................................................................................................................................... 223 16.2.1 Sample Collection and Preparation.................................................................................................................. 223 16.2.1.1 Sample Collection............................................................................................................................ 223 16.2.1.2 Preparation of the Food Matrix....................................................................................................... 223 16.2.2 Detection Procedures........................................................................................................................................ 224 16.3 Conclusions and Future Perspectives............................................................................................................................. 226 References.................................................................................................................................................................................. 226

16.1  Introduction 16.1.1  Classification of the Genus Micrococcus The genus Micrococcus was originally described by Cohn1 almost 140 years ago. After numerous reorganizations of the genus and reclassifications of certain species into new genera2,3 the genus now is encompassing a clade of four species including Micrococcus luteus4 the type species of the genus, Micrococcus lylae,4 Micrococcus antarcticus,5 and Micrococcus flavus.6 The species are separated from each other at 16S rRNA gene similarities between 96.9 and 98.3% but representatives of related species of the genus exhibit peptidoglycan type A2 (lysine based peptidoglycan and an interpeptide bridge consisting of the stem peptide) or A4α (lysine based peptidoglycan and aspartic acid as the interpeptide bridge) according to Schleifer and Kandler.7 Quinone systems consist predominantly of menaquinone with seven and/or eight isoprenoic units in the side chain and one of these units may be dihydrogenated such as MK-7 (H 2) and MK-8 (H2), MK-8 and MK-8 (H2) or MK-8 (H2). Chemotaxonomic characteristics such as peptidoglycan type and quinone system are considered to be highly conserved among species of a single genus with only few known exceptions. Already based on the characteristics so far collected for four species both quinone system and peptidoglycan type are unusually variable among species of the genus. The quinone system may be useful for identification of strains of M. flavus which has been reported to contain a quinone system consisting of the major compounds MK-7 (H 2) and MK-8 (H2). On the other hand M. lylae and M. antarcticus are characterized by quinone systems also found in certain M. luteus strains (Table 16.1) and hence they cannot

be identified based on this trait alone. On the other hand, peptidoglycan type A2 may be indicative for strains of M. luteus biovar I. Fatty acid profiles predominantly show iso-C15:0 and anteiso-C15:0 acids.4–6 Other fatty acids were reported only in minor amounts. Differences useful for differentiation between Micrococcus species are not reported. Hence, also fatty acid analysis, broadly applied in clinical diagnostics for rapid identification of pathogens apparently is not useful for reliable identification of Micrococcus species and especially for identification of M. luteus and M. lylae. Variability in physiological traits among strains of M. luteus4 makes it almost impossible to reliably distinguish this species from the other species of the genus. However, optimal growth temperatures suggest that M. luteus and M. lylae may possess the potential to cause human infections (Table 16.1). Among established bacterial species M. luteus is quite unique. Strains of this species exhibit significant variability in physiological traits as mentioned above but also in both peptidoglycan type and quinone system. Strains of this species either contain a peptidoglycan type A2 or A4α and the quinone system either consists of two major compounds MK-8 and MK-8 (H2) or only MK-8 (H2).4 Mainly based on this variability the species was recently subdivided into three biovars. Interestingly, strains of one of these biovars, biovar II exhibit a unique band after REP-PCR which is lacking in representatives of biovar I and biovar III.4 M. luteus is an inhabitant of the human skin whereas M. lylae is only rarely isolated from this habitat.8 Interestingly, at EMBL gene bank more than 50 sequence entries can be found which are sharing 99.5% or even higher 16S rRNA gene similarity with the type strain of this species. In contrast, only one entry can be found in the gene bank which shares 99.5% 221

222

Molecular Detection of Foodborne Pathogens

Table 16.1 Phenotypic Differentiation Between Micrococcus Species Characteristic Optimal growth temperature (°C)

M. luteus

M. lylae

M. flavus

M. antarcticus

37

37

31

16.8

Nitrate reduction

–

–

–

+

Voges–Proskauer reaction

–

–

–

+

MK-8, MK-8 (H2) or MK-8 (H2)

MK-8(H2)

MK-8 (H2), MK-7 (H2)

A2 or A4α

A4α

A4α

MK-8, MK-8 (H2) Not analyzed

–

–

+

– + – +

– – – +

– – + –

Major quinone(s) Peptidoglycan type

Hydrolysis of: Tween 80

v

D-Mannose D-Maltose L-Malate D-Trehalose

+ v v v

Assimilation of:

Sources: Wieser, M. et al., Int. J. Syst. Evol. Microbiol., 52, 629, 2002; Liu, H. et al., Int. J. Syst. Evol. Microbiol., 50, 715, 2000, and Liu, X.-Y. et al., Int. J. Syst. Evol. Microbiol., 57, 66, 2007.

16S rRNA gene sequence similarity with the type strain of M. lylae whereas several entries can found designated M. lylae but sharing less than 99.5% similarity. In bacterial systematics 16S rRNA gene sequences are not accepted as a tool for species identification because cases are known where genomically and phenotypically clearly separated species share 99.9% 16S rRNA gene similarity.9 However, values of 99.5% or higher may be suitable for preliminary identification of these strains as members of M. luteus or at least a very closely but so far undescribed species.

16.1.2 Biology, Pathogenesis, and Medical Importance The family Micrococcaceae includes Gram-positive bacteria found in a variety of different habitats e.g., water, various soils, insects, skin of humans and most other mammals and as a consequence are found on a diversity of different food. Nonspecific isolation of Micrococcus strains from grain, fruits, seeds, plants, milk, cheese, fresh meat and fermented meat, and fish products is common in food microbiology.10–13 On food some members of this genus are commensals in other cases different strains function as fermenters and ripers but also as spoilers as well as foodborne pathogens mainly caused by their production of biogenic amines (BA).14,15 M. luteus as well as M. lylae are known to be involved in ripening of certain naturally ripened products e.g., cheese, fermented fish products (e.g., shiokara, cassava), and salami type sausages. M. luteus is used as a starter culture strain for fermented fish products and fermented dry sausages. Nevertheless, Micrococcus spp. will never represent the main flora in neither of these products, which are lactobacilli, pediococci, staphylococci, and for

certain products yeasts and molds. However, in some fermented products Micrococcus spp. are considered to be essential for the development of aroma and flavor.10–13,16–19 Microbial enzymes produced by Micrococcus spp. which are involved in the development of flavor and aroma are a salt resistant glutaminase, an amylase, enzymes to degrade L-methionine to volatile sulfur compounds. Enzymes for protein hydrolysis and especially enzymes for lipolytic activity to modify generated carbonyl compounds are essential for the development of flavor. These activities have been shown to be essential for the specific odor of long air dried cured sausages.20 The action of enzymes required for ripening of fermented foods can cause spoilage of food as well. Hence, Micrococcus spp. as halotolerant microorganisms (5–10% NaCl),4 have been found in different spoiled food stuff. Fresh meat, sea food and insufficient thermal processing of products rich in fat are at risk to contain high numbers of Micrococcus spp. especially in the absence of other microbial spoilers as are pseudomonads and lactobacilli. Nevertheless, spoilage by Micrococcus spp. is rare. Typical changes include protein hydrolysis and lipolysis.21,22 Probably the greatest health threat for humans is related to the capability of micrococci to produce BA. As Micrococcus spp. are found on different raw, fermented, and sometimes even in heat treated foods many studies include Micrococcus as reference strains to test the effect of antimicrobial activity of different products or treatments or to study the effect of transfer from one environment to another which might be important for extraintestinal infections.23,24 BA are formed by decarboxylation of free amino acids. Many different bacterial species found in food are able to produce sufficient amounts of decarboxylase to be released

223

Micrococcus

in the food matrix and produce BA such as histamine, putrescine, cadaverine, tyramine, phenylethylamine, and further.25–27 Certain strains of M. luteus have also been shown to produce cadaverine in foods but there is no evidence from literature of capability for production of other BA.27 Cadaverine itself has only little toxic effects but it potentiates the toxicity of histamine in food by inhibiting histamine-metabolizing enzymes such as diamine oxidase and histamine N-methyl-transferase.28 The toxicity of histamine and other BA in humans are based on the interaction of histamine with cellular membrane receptors, effecting the cardiovascular system, smooth ­muscles, and eventually motor neuron stimulation. Symptoms in humans of biogenic amine poisoning are urticaria, flushing, headache, abdominal cramps, diarrhea, and vomiting. The effect varies individually (histamine intolerance) and it is dose and matrix dependent. Thus, the same amount of BA may cause no to severe symptoms if administered with different food matrices.25,29 The whole family of Micrococcus strains may also play an important role as normal gut microflora and may also be involved in symptoms of digestive disorders in humans.30 Thus, the role of Micrococcus spp. in food is highly diverse as the genus itself. Additionally, strains of the genus are known as opportunistic pathogens for extraintestinal disease mainly in immunocompromised persons,31,32 meningitis,33 septic arthritis,34 endocarditis,35–37 infections of leukemia patients with indwelling lines and patients undergoing continuous amulatory peritoneal dialysis or with ventriculo-peritoneal shunt,38 intracranial suppuration,39 bacteraemia,40,41 chronic cutaneous infections in HIV positive patients,42 and catheter infection.43

16.1.3 Identification and Diagnosis of the Genus Micrococcus For the sampling of food and food stuff—if no specific advice is given—the specific International Standard has to be followed dealing with the product concerned e.g., DIN 10162. A portion of the product is homogenized with 1:10 of adequate dilution medium e.g., peptone solution (ISO 6887) using a stomacher 400 or by Ultra turrax. A dilution series (depending of expected colony counts) are plated. For meat and meat products, cheese as well as for fish and fish products in which Micrococcus spp. may be found the detection and isolation of Micrococcus spp. is difficult as most strains are not able to compete with Staphylococcus spp.16,20 In most cases Micrococcus spp. are not able to grow on Baird–Parker agar, incubated aerobically at 30 or 37°C for 48 h and thus, cannot be isolated in parallel with Staphylococcus spp. even though, some articles and books refer to it.44 In some cases M. luteus and M. lylae can be detected on colony count agar especially if the product harbors a low number of bacteria. Suggestions for the isolation of M. luteus with the addition of 20 µg/ml furazolidone to colony count agar are confirmed in our studies.45,46 However, since the existence of furazolidone sensitive micrococci was reported47 not all members of the

micrococci community will be detected using this selective medium. In products containing a rather high number of other bacterial species a combination of 5% sodium chloride and 20 µg/ml furazolidone has been found useful.46 Furthermore, verification of colonies by aerobic/anaerobic growth to distinguish between facultative anaerobes and Gram staining has to be done. Discrimination from e.g., Kocuria varians and differentiation between the Micrococcus species by biochemical reactions such as tests for katalase reaction, lysostaphin resistance testing, assimilation of lactose, glucose, gelatin, glycerol, and nitrate reduction (see Table 16.1) is complicated by highly similar reaction profiles. Thus, there is an essential need for molecular tools allowing identification of members of the genus Micrococcus as well as to discriminate between species. To the best of our knowledge no molecular detection method for Micrococcus from food has been developed.

16.2  Methods 16.2.1  Sample Collection and Preparation 16.2.1.1  Sample Collection As M. luteus and M. lylae are found in a variety of different food with unpredictable amounts ranging from one to 105 cfu per gram/ml, respectively, the sampling procedure and preparation are dependent on the product as well on the purpose of detection. Hence, not only general procedures have to be followed, such as cooling and fast transport to the laboratory as well as rapid sample analysis but also to control these specific food products at different processing steps, at different times of fermentation, or during shelf life. Further chemical analysis on the food matrix might be necessary e.g., analysis of BA to define safety, accurate shelf life, or consistent sensorial quality. In some cases it might be necessary to analyze Micrococcus isolates for their ability to cause biogenic amine production in food, to define their spectrum of lipolytic activity, protein hydrolysis and/or their carbohydrate degradation ability to categorize the isolate as spoiler, fermenter or pathogen. However, it appears to be feasible to replace some of these growth dependent examinations by development of PCRs directed to genes coding for certain enzymes such as ornithine- and lysine-decarboxylase or arginine-dihydrolase. 16.2.1.2  Preparation of the Food Matrix The complexity of food matrices, their preparation and the choice of the best possible way to detect different bacteria or elute bacterial DNA are still considered as one of the upper most important problems in food microbiology and thus, have been described, experimentally processed and extensively discussed for most foodborne pathogens,48,49 but so far not for Micrococcus spp. Nevertheless, one can assume that protocols for preparation of food matrices used for the isolation and identification of other foodborne pathogens might be useful also for micrococci.

224

Meat and meat products. Micrococcus spp. occurs in fermented meat products as well as products conserved by high salt concentrations and low temperature thermal processing. Examples of these products are dry-fermented sausages, traditional made and temperature sensitive products like pâté containing liver or lard.11,12,18,19 These products are difficult to process further for microbial DNA analysis as they contain substances e.g., proteinases that are able to disturb microbial DNA extraction as well as DNA polymerase activity for amplification.49,50 Additionally the removal of fat, proteins as well as DNA from meat species is challenging. Nonetheless, there are methods that are considered to allow direct DNA isolation from meat and meat products and further amplification e.g., real-time PCR and thus, quantification of microorganisms.50–53 In the case of dry fermented sausage as well as products containing lard, the separation of fat is most important. Various filtration and centrifugation steps prior to DNA preparation are essential.51,53 Dairy products. Micrococcus spp. are reported to be found on smear ripened cheese e.g., Gubeen cheese on traditionally made goat and sheep cheese, and on Camembert-like and blue-veined cheeses also.16,17 In some cases it is not essential to homogenize cheese if only the surface flora needs to be analyzed.54 But to analyze total cheese one has to homogenize a specific volume of the dairy product in adequate suspension e.g., 0.1% peptone solution or physiological sodium chloride solution. Like certain meat products dairy food matrices may harbor substances to reduce the amount of extracted DNA and/or inhibit further DNA amplification. Fish and seafood. Micrococcus spp. are reported to be mainly found in traditionally fermented fish (e.g., cassava fish for further production of lanhouin) or fish sausage.17,18 As fish and especially fish products are at risk to contain significant high values of BA, the detection of Micrococcus spp. which are able to produce BA in fish and sea food is relevant to human health.25 Thus, next to the isolation and/ or detection of Micrococcus spp. it is important to identify their ability to produce BA by molecular or traditional based methods. A suitable and easy isolation method for Micrococcus DNA extraction from all above mentioned food matrices is given by Stevens and Jaykus55 and by Rossmanith et al.56 Both methods are described below and are slightly adapted to fit the purpose: (1) Eleven grams of the sample is diluted in 25 ml sterile 0.9% saline solution and 8 ml of 25% w/v sodium citrate solution supplemented with polyethylene glycol to a final concentration of 4% and treated by a stomacher for at least 2 min at room temperature. For meat and fish products containing larger particle the homogenized product is filtered through cheesecloth. Samples are centrifuged at 9700×g for 15 min at 4°C. The pellet is transferred to DNA extraction using either DNAzol or a commercial kit for total microbial DNA extraction.

Molecular Detection of Foodborne Pathogens

(2) The sample is diluted 1:2 in lysis buffer containing 1× PBS, 8 M urea and 1% SDS. The sample is treated by a stomacher for 3 min. Lysis buffer is added to a volume of 40 ml and lysis is carried out at 45°C for 30 min at constant shaking at 200 rpm following a centrifugation step for 30 min at room temperature and 3220×g. The supernatant is discharged and the pellet dissolved in 500–1500 µl of 1× PBS. Hard cheese and sausage samples are sedimentated for 5 min and the supernatant is further centrifuged at 8000×g for 5 min. DNA isolation from the remaining pellet is performed using a commercial kit for total microbial DNA isolation.

16.2.2  Detection Procedures (i) PCR detection of M. luteus: So far there are no standard protocols used for detection and identification of Micrococcus spp. on a molecular level but the following procedure can be used for obtaining template DNA for proposed PCR amplification (see Table 16.2). Suspective colonies on e.g., LB agar containing 5% sodium chloride and 20 µg/ml furazolidone can be directly used for template DNA extraction as described by three cycles of freeze-thawing.57 (1) One loop of biomass is suspended in 50–100 µl sterile ddH20. (2) rapidly frozen in liquid nitrogen. (3) followed by melting at 60°C for 5min. (4) Repeat steps 2 and 3 twice. (5) Cell debris is pelleted in a microcentrifuge at approximately 10000×g. The supernatant is ready for use in PCR (and can be stored at –20°C for several weeks). If colony counts are expected at 104 per g/ml of food a direct molecular detection might be possible and total microbial DNA isolation can be performed as described in Section 16.2.2. PCR and sequencing of the 16S rRNA gene might be performed as follows:



(1) Quantification of template DNA. (2) Use 5 ng (extracted from culture). (3) 10× dNTPs (2 mM each). (4) 100 mM forward primer and 100 mM reverse primer (stock) (see Table 16.3). (5) PCR reaction buffer including MgCl2 (10×). (6) Taq polymerase (e.g., 5 U/µl). (7) A typical reaction of 25 µl includes: 1–5 µl template DNA, 2.5 µl 10× dNTPs, 0.1 µl forward primer, 0.1 µl reverse primer, 2.5 µl 10× PCR buffer, 0.5 U Taq polymerase (0.1µl of 5 U/µl) adjust with ddH2O to 25 µl. (8) PCR conditions are given in Table 16.3.

225

Micrococcus

Table 16.2 Primers and PCR Conditions Primer Com1 Com2 ML-recA-for ML-recA-rev

Sequence (5′–3′) cagcagccgcggtaatac ccgtcaattcctttgagttt gccctggacccggtctacgcccg cgccgatcttctcgcgcagctgg

Target Gene 16S RNA gene variable region V4-5 recA

Table 16.3 Characteristics Differentiating Micrococcus luteus and Kocuria varians (formerly Micrococcus varians) Characteristics Aerobic acid from glucose Nitrate reduced to nitrite Simmons citrate agar Inorganic nitrogen agar

Micrococcus luteus

Kocuria varians

– – – +

+ + + –

Sources: Kocur, M., In Bergey’s Manual of Systematic Bacteriology, Sneath, P.H.A. et al. (eds.), Williams & Wilkins, Baltimore, 1986.

(9) Amplicons can be purified directly or after detection in gel electrophoreses and excised from the matrix using different commercial kits. (10) Subject to automated sequencing using the above mentioned forward and/or reverse primers, match, and analyze sequence to existing database. Construction of specific primers for the detection of micrococci is hampered by the limited genomic sequences deposited at gene banks. The 16S rRNA gene is most likely not suitable to reach this goal because of the highly conserved character of this gene. More promising might be primers directed to so-called house-keeping genes which are less conserved then 16S rRNA genes but sufficiently conserved to construct Micrococcus-specific primers. However, among commonly studied house-keeping genes we could only find nine recA gene sequences of micrococci in gene banks of which only two were of well characterized strains, M. luteus ATCC 381 (AF214783) and Micrococcus lylae ATCC 27566 (AF214778). After alignment with corresponding gene of two Arthrobacter spp. (closely related to Micrococcus) we were able to construct a primer pair (see Table 16.3) which should allow specific amplification of partial recA with M. luteus and M. lylae resulting in an amplicon of 310 bp. However the suitability, selectivity, and specificity of this approach have not been tested so far because of unavailability of sufficient number of well characterized strains of both species. (ii) Molecular detection of biogenic amine ­producing bacteria: Some Gram-positive species (e.g., Clostri­dium, Micrococcus, Lactobacillus, Leuconostoc, and Tetragen­ ococcus) generate BA including histamine, tyramine,

Amplicon Size (bp) variable 310

Annealing Temperature

Reference

54°C

Schwieger et al.58 This study

–

phenylalanine, putrescine, and cadaverine from free amino acids in fermented foods and beverages as a result of their amino acid decarboxylase activities. Although BA such as histamine and tyramine do not affect the sensory quality of the food products, these compounds have been implicated in a number of food poisoning cases. These bacteria harbor genes hdc and tyrdc encoding for histidine and tyrosine decarboxylases, respectively, which are involved in the production of BA from free amino acids. By employing primers targeting hdc and tyrdc genes as well as 16S rRNA gene of bacteria as an internal PCR control, Coton, and Coton60 developed a multiplex PCR for the early detection of potential tyramine and histamine producing bacteria directly on bacterial colonies. Specifically, the following three primer sets are utilized: HDC3 (5′-GATGGTATTGTTTCKTATGA-3′) and HDC4 (5′-CAAACACCAGCATCTTC-3′) for a 440-bp hdc gene fragment; TD2 (5′-ACATAGTCAACCATRTTGAA-3′) and TD5 (5′-CAAATGGAAGAAGAAGTAGG-3′) for a 1100-bp tyrdc gene fragments; and 16S rRNA universal primers BSF8 (5′-AGAGTTTGATCCTGGCTCAG-3′)  a nd  BSR15 41 (5′-AAGGAGGTGATCCAGCCGCA-3′) for a 1530-bp fragment as the PCR internal control. Primer concentrations are 20 pmol for HDC3, HDC4, TD2, and TD5 and 5 pmol for BSF8 and BSR1541; and the PCR amplification program consists of one cycle of 95°C for 5 min; 32 cycles of 95°C for 45 sec, 52°C for 45 sec, and 72°C 1 min 15 sec; and one cycle of 72°C for 5 min. After completion, 18 μl of each PCR sample are separated on 0.8% (wt/vol) agarose gels in 1× TBE buffer at 100 V for 50 min and then visualized with ethidium bromide staining using a GelDoc 2000. The authors successfully detected HDC+/TyrDC+ strains of Clostridium, Enterococcus, Lactobacillus, Oenococcus, and Tetragenococcus. Unfortunately, while pyruvoyl-dependent HDC activity has been observed in the genus Micrococcus, the hdc, and tyrdc-equivalent gene sequences are not found in micrococci, which prevent the PCR application of the HDC3, HDC4, TD2, and TD5 primers for the detection of tyramine and histamine producing micrococci. Perhaps, the gene responsible for pyruvoyl-dependent HDC activity in micrococci may have evolved over time so that it differs significantly from that in Clostridium, Enterococcus, Lactobacillus, Oenococcus, and Tetragenococcus. Further examination of micrococcal genome is clearly needed to identify distinct gene sequence accounting for its pyruvoyldependent HDC activity for subsequent PCR application.

226

16.3 Conclusions and FUture Perspectives In food microbiology species of the genus Micrococcaceae have been described as fermenters, spoilers, and pathogens. But because they do not attach major importance in one of these multifarious functions not much research has been undertaken to clarify their role in food. In most cases isolation of micrococci is mainly coincidently as no standard protocols exist in food microbiology for isolation or even further characterization. The recent reclassifications of certain species especially the classification of e.g., M. varians into K. varians sets new requirements in differentiation of micrococci. The differentiation between M. luteus and K. varians is essential to studying fermentation and spoilage in food microbiology, as they have different functions in food fermentation, production of BA and/or spoilage. For instance K. varians is known to be a beneficial microorganisms of main importance in food fermentation and interaction with other fermenters e.g., staphylococci.61 Nevertheless, the differentiation to micrococci will cause problems because both species have similar growth requirements and hence, are difficult to discriminate (see Table 16.3). Therefore, new methods have to be described, adopted, and evaluated for detecting this genus in food. In order to develop a PCR system for rapid identification two strategies appear to be most promising: (i) Development of a PCR which generates genomic fingerprints characteristic for Micrococcus species. Recently, it has been demonstrated that M. luteus biovar II strains exhibit a characteristic band after REP-PCR which was not detected in representatives of biovar I and III strains. This observation indicates a rather high genomic variability among strains of this species which is also demonstrated by the high degree of variability in phenotypic characteristics and it may be difficult to construct primers which in PCR produce M. luteus-specific fingerprints. (ii) Development of a species specific-PCR. In order to develop such an approach sequence information from house-keeping genes are urgently needed. These genes should be present in all Micrococcus species and strains and sufficiently variable to construct primers for specific identification at the species level. Unfortunately, quite a few sequences other than 16S rRNA gene sequences are available from gene banks which hamper the design of such a PCR-approach. Since the biggest threat for humans is the capability of M. luteus to decarborxylate lysine and to produce the biogenic amine cadaverine the most useful tool would be a PCR specifically detecting the gene coding for lysine-decarboxylase but also for this gene no sequence information is available concerning M. luteus. So far, it is unknown whether lysine decarboxylation activity is a common trait of M. luteus or only certain strains of this species and hence, such a PCR would have the advantage to discriminate between hazardous and nonhazardous strains of the species. However, also for this gene no sequence information from M. luteus is accessible. Hopefully, in the near future the complete genome of a representative of a Micrococcus

Molecular Detection of Foodborne Pathogens

species will be sequenced. This information would be a huge step forward to identify house-keeping genes or other gene target and their sequences and to construct species- or genusspecific primers. However, for any of these developments a large number of reliable identified strains of the species in question will have to be subjected to the system to test its suitability. The two species M. luteus and K. varians share in the the 16S rRNA genes less than 95% similarity and hence a PCR may be developed to distinguish the two species. Since the 16S rRNA similarities are quite low it appears to be achievable to develop a duplex PCR with three or four primers which amplifies two 16S rRNA gene fragments of different sizes specific for M. luteus and K. varians, respectively.

References













1. Cohn, F. Untersuchungen über Bakterien. Beitr. Biol. Pfanz., 1, 127, 1872. 2. Stackebrandt, E. et al. Taxonomic dissection of the genus Micrococcus: Kocuria gen. nov., Nesterenkonia gen. nov., Kytococcus gen. nov., Dermacoccus gen. nov., and Micrococcus Cohn 1872 gen. emend. Int. J. Syst. Bacteriol., 45, 682, 1995. 3. Koch, C., Schumann, P. and Stackebrandt, E. Reclassification of Micrococcus agilis (Ali-Cohen 1889) to the genus Arthrobacter as Arthrobacter agilis comb. nov. and remendation of the genus Arthrobacter. Int. J. Syst. Bacteriol., 45, 837, 1995. 4. Wieser, M. et al. Emended descriptions of the genus Micrococcus, Micrococcus luteus (Cohn 1872) and Micrococcus lylae (Kloos et al. 1974). Int. J. Syst. Evol. Microbiol., 52, 629, 2002. 5. Liu, H. et al. Characterization of Micrococcus antarcticus sp. nov., a psychrophilic bacterium from Antarctica. Int. J. Syst. Evol. Microbiol., 50, 715, 2000. 6. Liu, X.-Y. et al. Micrococcus flavus sp. nov., isolated from activated sludge in a bioreactor. Int. J. Syst. Evol. Microbiol., 57, 66, 2007. 7. Schleifer, K.H. and Kandler, O. Peptidoglycan types of bacterial cell walls and their taxonomic implications. Bacteriol. Rev., 36, 407, 1972. 8. Kloos, W.E. and Musselwhite, M.S. Distribution and persistence of Staphylococcus and Micrococcus species and other aerobic bacteria on human skin. Appl Microbiol., 30, 381, 1975. 9. Stackebrandt, E. and Goebel, B.M. 1 Taxonomic note: a place for DNA-DNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology. Int. J. Syst. Bacteriol., 44, 846, 1994. 10. Anihouvi, V.B. Microbiological changes in naturally fermented cassava fish (Pseudotolithus sp.) for lanhouin production. Int. J. Food Microbiol., 116, 287, 2007. 11. Coppola, R. et al. Characterization of micrococci and ­staphylococci isolated from soppressata molisana, a Southern Italy fermented sausage. Food Microbiol., 14, 47, 1997. 12. Garcia Fontán, M.C. et al. Microbiological characteristics of “androlla”, a Spanish traditional pork sausage. Food Microbiol., 24, 52, 2007. 13. Gardini, F., Tofalo, R. and Suzzi, G. A survey of antibiotic resistance in Micrococcaceae isolated from Italian dry fermented sausages. J. Food Prot., 6, 937, 2003.

Micrococcus 14. Prado, B. et al. Numerical taxonomy of microorganisms isolated from goat cheese made in Chile. Curr. Microbiol., 43, 396, 2001. 15. Straub, B.W. et al. The formation of biogenic amines by fermentation organisms. Z. Lebensm. Unters. Forsch., 201, 79, 1995. 16. Fernandez-García, E., Tomillo, J. and Nuñez, M. Formation of biogenic amines in raw milk Hispánico cheese manufactured with proteinases and different levels of starter culture. J. Food Prot., 63, 1551, 2000. 17. Addis E. et al. The growth, properties and interactions of yeasts and bacteria associated with the maturation of Camembert and blue-veined cheeses. Int. J. Food Microbiol., 69, 25, 2001. 18. Aryanta, R.W., Fleet, G.H. and Buckle, K.A. The occurrence and growth of microorganisms during the fermentation of fish sausage. Int. J. Food Microbiol., 13, 143, 1991. 19. Cordero, M.R. and Zumalacárregui, J.M. Characterization of micrococcaceae isolated from salt used for Spanish dry-cured ham. Lett. Appl. Microbiol., 31, 303, 2000. 20. Vilar, I. et al. A survey on the microbiological changes during the manufacture of dry-cured lacón, a Spanish traditional meat product. J. Appl. Microbiol., 89, 1018, 2000. 21. Lücke, F.-K. Fermented sausages. In: Microbiology of Fermented Foods. 2nd Edn. Vol 2. pp. 441–483. B.J.B. Wood (ed.). Blackie Academic & Professional, London, United Kingdom, 1998. 22. Jay, J.M. A review of recent taxonomic changes in seven genera of bacteria commonly found in foods. J. Food Prot., 66, 1304, 2003. 23. Sierra, M. et al. Numerical taxonomy of an ‘atypical’ population of gram-positive cocci isolated from freshly dressed lamb carcasses. Int. J. Food Microbiol., 24, 363, 1995. 24. Mauriello, G. et al. Antimicrobial activity of a nisin-activated plastic film for food packaging. Lett. Appl. Microbiol. 41, 464, 2005. 25. Lehane, L. and Olley, J. Histamine fish poisoning revisited. Int. J. Food Microbiol., 58, 1, 2000. 26. Leuschner, R.G., Heidel, M. and Hammes, W.P. Histamine and tyramine degradation by food fermenting microorganisms. Int. J. Food Microbiol., 39, 1, 1998. 27. Lakshmanan, R., Shakila, R.J. and Jeyasekaran, G. Changes in the halophilic amine forming bacterial flora during saltdrying of sardines (Sardinella gibbosa). Food Res. Int., 35, 541, 2002. 28. Taylor, S.L. and Sumer, S.S., Determination of histamine, cadaverine and putrescine. In: Seafood Quality Determination. Proceedings of an International Symposium. pp. 245–253. Kramer, D.E. and Liston, J. (eds.). Elsevier Science, Amsterdam, 1986. 29. Bardocz, S. Polyamines in food and their consequences for food quality and human health. Trends Food. Sci. Technol., 6, 341, 1995. 30. Bouhnik, Y. et al. Bacterial populations contaminating the upper gut in patients with small intestinal bacterial overgrowth syndrome. Am J. Gastroenterol., 94, 1327, 1999. 31. Ronald, J. et al. Micrococcus-associated central venous catheter infection in patients with pulmonary arterial hypertension. Chest, 126, 90, 2004. 32. Albertson, D., Natsios, G.A., and Gleckman, R. Septic shock with Micrococcus luteus. Arch. Intern. Med., 138, 487, 1978. 33. Fosse, T. et al. Meningitis due to Micrococcus luteus infection. Infection, 13, 280, 1985.

227 34. Wharton, M. et al. Septic arthritis due to Micrococcus luteus. J. Rheumatol., 13, 659, 1986. 35. Glupezynski, Y. et al. Clinical evaluation of teicoplanin for therapy of severe infections caused by gram-positive bacteria. Antimicrob. Agents Chemother., 29, 52, 1986. 36. Dürst, U.N. et al. Micrococcus luteus: a rare pathogen of valve prosthesis endocarditis. Z. Kardiol., 80, 294, 1991. 37. Seifert, H., Kaltheuner, M. and Perdreau-Remington, F. Micrococcus luteus endocarditis: case report and review of the literature. Int. J. Med. Microbiol., 282, 431, 1995. 38. Magee, J.T. et al. Micrococcus and Stomatococcus spp. from human infections. J. Hosp. Infect., 16, 67, 1990. 39. Selladurai, B. et al. Intracranial suppuration caused by Micrococcus luteus. Br. J. Neurosurg., 7, 205, 1993. 40. von Eiff, C. et al. Micrococcus luteus as a cause of recurrent bacteremia. Pediatr. Infect. Dis. J., 15, 711, 1996. 41. Peces, R. et al. Relapsing bacteraemia due to Micrococcus luteus in a haemodialysis patient with a Perm-Cath catheter. Nephrol. Dial. Transplant., 12, 2428, 1997. 42. Smith, K.J. et al. Micrococcus folliculitis in HIV-1 disease. Br. J. Dermatol., 141, 558, 1999. 43. Oudiz, R.J. et al. Micrococcus-associated central venous catheter infection in patients with pulmonary arterial hypertension. Chest, 126, 90, 2004. 44. Gleman, A., Drabkin, V. and Glatman, L. Evaluation of lactic acid bacteria, isolated from lightly preserved fish products, as starter cultures for new fish-based food products. Innov. Food Sci. Emerg., 1, 219, 2000. 45. von Rheinbaben, K.E. and Hadlok, R.M. Rapid distinction between micrococci and staphylococci with furazolidone agars. Antonie Van Leeuwenhoek, 47, 41, 1981. 46. Muhri, S., Busse, J. and Hilbert, F., unpublished results. 47. Baker, J.S. Comparison of various methods for differentiation of staphylococci and micrococci. J. Clin. Microbiol., 19, 875, 1984. 48. Powell, H.A. et al. Proteinase inhibition of the detection of Listeria monocytogenes in milk using polymerase chain reaction. Lett. Appl. Microbiol., 18, 59, 1994. 49. Wilson, I.G. Inhibition and facilitation of nucleic acid amplification. Appl. Environ. Microbiol., 63, 3741, 1997. 50. Taylor, T.M. et al. Effect of food matrix and cell growth on PCR-based detection of Escherichia coli O157:H7 in ground beef. J. Food Prot., 68, 225, 2005. 51. Fukushima H. et al. Rapid separation and concentration of food-borne pathogens in food samples prior to quantification by viable-cell counting and real-time PCR. Appl. Environ. Microbiol., 73, 92, 2007. 52. Rodríguez-Lázaro, D. et al. Rapid quantitative detection of Listeria monocytogenes in salmon products: evaluation of pre-real-time PCR strategies. J. Food Prot., 68, 1467, 2005. 53. Stevens, K.A. and Jaykus, L.A., Bacterial separation and concentration from complex sample matrices: a review, Crit. Rev. Microbiol., 30, 7, 2004a. 54. Brennan, N.M. et al. Biodiversity of the bacterial flora on the surface of a smear cheese. Appl. Environ. Microbiol., 68, 820, 2002. 55. Stevens, K.A. and Jaykus, L.A. Bacterial separation and ­concentration from complex sample matrices: a review. Crit. Rev. Microbiol., 30, 7, 2004. 56. Rossmanith, P. et al. Development of matrix lysis for concentration of gram positive bacteria from food and blood. J. Microbiol. Methods, 69, 504, 2007. 57. Wieser, M. and Busse, H.-J. Rapid identification of Staphylococcus epidermidis. Int. J. Syst. Evol. Microbiol., 50, 1087, 2000.

228 58. Schwieger, F. and Tebbe, C.C. A new approach to utilize PCR-single-strand-conformation polymorphism for 16S rRNA gene-based microbial community analysis. Appl. Environ. Microbiol., 64, 4870, 1989. 59. Kocur, M., Genus I. Micrococcaceae Cohn 1872, 151AL. In Bergey’s Manual of Systematic Bacteriology. Vol. 2, pp. 1004–1008. Sneath, P.H.A. et al. (eds.). Williams & Wilkins, Baltimore, 1986.

Molecular Detection of Foodborne Pathogens 60. Coton, E. and Coton, M. Multiplex PCR for colony direct detection of Gram-positive histamine- and tyramine-producing bacteria. J. Microbiol. Methods, 63, 296, 2005. 61. Tremente, P. et.al. Interaction of Staphylococcus cylosus and Kocunia Varians isolated from fermented meats. J. Apple. Microbiol., 103, 743, 2007.

17 Mycobacterium Irene R. Grant

Queen’s University Belfast

Catherine E.D. Rees

University of Nottingham

Contents 17.1 Introduction ................................................................................................................................................................... 229 17.1.1 Classification of Mycobacterium avium subsp. paratuberculosis ................................................................... 229 17.1.2 Biology, Pathogenesis, and Medical Importance.............................................................................................. 230 17.1.3 Current Methods for the Detection of Foodborne Map.................................................................................... 231 17.1.3.1 Phenotypic Techniques.................................................................................................................... 231 17.1.3.2 Bacteriophage-Based Techniques.................................................................................................... 232 17.1.3.3 Molecular Techniques...................................................................................................................... 232 17.2 Methods.......................................................................................................................................................................... 237 17.2.1 Reagents and Equipment................................................................................................................................... 237 17.2.2 Sample Preparation .......................................................................................................................................... 237 17.2.3 Detection Procedures . ..................................................................................................................................... 238 17.3 Conclusions and Future Perspectives ............................................................................................................................ 239 References.................................................................................................................................................................................. 240

17.1  Introduction 17.1.1 Classification of Mycobacterium avium subsp. paratuberculosis The genus Mycobacterium in the family Mycobacteriaceae contains both fast-growing and slowgrowing species.1 The slow-growing species include the important animal or human pathogenic species Mycobacterium tuberculosis (M.tb), Mycobacterium bovis, Mycobacterium leprae and Mycobacterium avium subsp. avium. Being most closely related to M. avium subsp. avium and M. avium subsp. silvaticum,2 M. avium subsp. paratuberculosis (Map) is an acid-fast, rod-shaped bacterium possessing a thick, lipidrich (waxy) cell wall, which confers on it the properties of acid-fastness (the ability to resist decolorisation by acidified alcohol in the Ziehl-Neelsen stain) and hydrophobicity. The lipid-rich cell wall gives mycobacteria, generally, a survival advantage and increased resistance to chemicals (e.g., chlorine3) and physical processes (e.g., pasteurization,4,5 UV light6), but the consequence of this defense is slow growth due to restricted uptake of nutrients through the cell wall. Characteristics that distinguish Map from other Mycobacterium spp. include its extremely slow growth (up to 18 weeks to form colonies on solid media) and its inability to produce mycobactin (an ­iron-chelating compound). This

requirement for culture media to be ­supplemented with synthetic mycobactin J for growth to occur in vitro is used as a differential diagnostic for the species. A conundrum exists however as under appropriate conditions in vivo the organism must be capable of much more rapid growth; this is demonstrated by the fact that high numbers are found in the feces of cattle showing clinical symptoms of disease despite digesta flow rates in the range of 3–5 liters per hour (see Croom et al.7). Molecular identification relies on the detection of specific signature sequences which may be either insertion elements which tend to be present in multicopies (IS900, ISMav2, ISMap02; see Section 17.1.3.3 (i)) or single copy genes within the genome of Map (F57, hspX, locus 251 and locus 255; see Table 17.1 and later discussion of specificity of these different Map-specific targets). Sequencing of the complete genome of Map strain K-10 was carried out by researchers at the University of Minnesota, in collaboration with the National Animal Disease Center, Ames, Iowa. The Map genome sequence was released in January 2004 and deposited in the GenBank database (accession no. AE016958). Analysis showed that Map K-10 has a single circular sequence of 4,829,781 base pairs, with a G + C content of  69.3%. A comparison of the genome sequences of Map and three other Mycobacterium spp. (M.

229

230

Molecular Detection of Foodborne Pathogens

Table 17.1 Map-Specific PCR Targets (in Chronological Order of Publication) Target

GenBank Accession No.

Type of Target

No. of Copies per Map Cell

Reference

14–20 1 1

51 59 60

≥3 1

61

6

63

IS900 F57 hspX

X16293 X70277

Insertion element Gene locus Gene

ISMav2

AF286339

Insertion sequence

Loci 251 and 255 ISMap02

Genes EF514846

Insertion element

tuberculosis, M. leprae and M.  bovis) was recently published and this has shed some new light on the genetic basis of mycobactin J dependence and slow growth in Map,8 as well as assisting in the continuing search for novel Mapspecific sequences for use in PCR-based detection methods and the development of microarray analysis methods (see Section 17.1.3.3 (v)).

17.1.2 Biology, Pathogenesis, and Medical Importance Map is well known as an animal pathogen, being the causative agent of Johne’s disease in ruminants,9 but other animals, such as alpacas, rabbits, foxes, stoats, and weasels can also be infected. Johne’s disease is prevalent in domestic animals worldwide (cattle, sheep, and goats principally) and has significant economic impacts for producers. It is generally believed that calves become infected by Map early in life by ingestion of the bacterium in infected milk or colostrum, or via an infected farm environment (fecal-oral route), and it takes a few years (typically 2–5 years) for clinical symptoms of Johne’s disease (weight loss, emaciation, chronic persistent diarrhea, failure to thrive) to become evident. Map is known to be shed in the milk and feces of infected animals. Infections with Map can be latent for years and diagnosis of paratuberculosis can be a challenge in the latent stages of the infection. ELISA tests are used to identify Map-infected animals and an interferon-γ assay (IFN-γ) to identify those where the infection has progressed to the disseminated state. For a recent review of the efficacies of these immunological tests see Nielsen and Toft.10 Once animals progress to the clinical stage of infection, feces may contain up to 1010 Map/g whereas milk aseptically obtained from the udder of an infected animal is thought to contain low levels of Map (2–8 CFU per 50 ml11). However, fecal contamination can, and does, occur during the milking process so low natural levels of Map in a cow’s milk may be augmented by Map from feces. Unfortunately, current cultural methods for Map do not permit accurate enumeration to verify the numbers of viable Map cells present in raw milk. A possible link between Map and Crohn’s disease in humans—though not necessarily causative—has long been suspected but this remains a contentious issue. Crohn’s

62

disease is a chronic inflammatory bowel disease of unknown etiology that typically affects young people in the 15–25 year age group. The clinical and pathological resemblance of Crohn’s disease and Johne’s disease was first recognized as long ago as 1913. The two diseases share common symptoms (chronic weight loss and diarrhea) and common pathology (ileum most frequently involved and granulomatous inflammation),12,13 but there are also distinct differences, for example acid-fast bacteria are not visible in gut sections from Crohn’s patients whereas abundant acid-fast bacteria are visible in sections of gut from cattle affected by Johne’s disease. Opinion in the gastroenterology community remains divided about the role, if any, of Map in Crohn’s disease. The current thinking regarding the aetiology of Crohn’s disease is that it is multifactoral with genetic predisposition (NOD2/CARD15 mutations on chromosome 16, along with the recently identified autophagy genes ATG16L1 and IRGM14), exogenous factors (possibly an infectious agent such as Map) and host specific-factors (e.g., “leaky gut,” vascular supply, hormones, neuronal activity) acting together to induce a chronic state of dysregulated mucosal immune function.15 A number of systematic reviews and meta-analyses of the evidence linking Map to Crohn’s disease have been published recently.16–18 Feller et al.16 considered both PCR and ELISA studies, Abubakar et al.17 considered only results of studies involving nucleic-acid based methods, whilst Waddell et al.18 considered all methods used to detect Map in Crohn’s patients (culture, PCR, FISH, DNA hybridization, and ELISA). The consensus opinion of these three studies, a recent review by Behr and Kapur,19 and indeed previous reviews of the evidence linking Map and Crohn’s disease by the UK Food Standards Agency, European Commission, Food Safety Authority of Ireland and UK National Association for Crohn’s and Colitis (previously summarized by Grant20), is that there is evidence of an association between Map and Crohn’s disease. Feller et al.16 concluded that the association seems to be specific. Waddell et al.18 concluded that evidence for the zoonotic potential of Map is not strong, but should not be ignored. The outstanding unanswered question highlighted by all reviews is: what is Map’s role in the etiology of Crohn’s disease? Further research is necessary on the medical front to answer this question. Assuming Map does have a role in Crohn’s disease and is, therefore, zoonotic, the most likely candidates as vehicles

231

Mycobacterium

of transmission of Map from animals to humans are milk and other dairy products, ground beef and water. Readers are directed to comprehensive reviews of the evidence of Map in food by Gould et al.,21, Grant22, and Griffiths.23 Cows’ milk has received most attention to date because, as described above, ingestion of Map-infected colostrum or milk is recognized as a primary means of transmission of Johne’s disease from cow to calf,24 and to control the spread of Johne’s disease within a herd it is recommended that calves are not fed such milk.25 Cows’ milk forms a significant element of the diet of most people, particularly that of young children, and therefore transmission of Map infection to vulnerable groups via milk is plausible. As most milk is consumed after pasteurization the focus of much of the research on Map in milk has been to assess the efficacy of milk pasteurization to inactivate Map. It is becoming more widely accepted that, despite the fact that HTST pasteurization at minimum 72oC for at least 15 sec should, according to numerous pasteurization experiments (reviewed by Grant22), theoretically achieve a 3–7 log10 reduction in numbers of this bacterium, that low numbers of viable Map may be present in commercially HTST pasteurized milk on occasion. The findings of several large scale surveys of retail pasteurized milk in the UK, Republic of Ireland, Czech Republic, USA, and Argentina illustrate this fact (reviewed by Grant20). The mechanism of this low-level Map survival has yet to be elucidated but estimates of the number of Map surviving HTST pasteurization are: 4–20 CFU/50 ml,26 10–20 CFU/150 ml,27 and 0.002–0.004 CFU/ ml.28 The public health consequences of low levels of Map in pasteurized cows’ milk being periodically consumed by susceptible individuals remains uncertain. Foods other than cow’s milk have not been extensively studied to date. Johne’s disease also affects sheep and goats and Map has been detected and/or isolated from the milk of these domesticated animals,29–31 and cheese made from these types of milk.32 Theoretically beef could also be a potential food vehicle of transmission of Map to humans, although to date there is no evidence of Map being detected in retail ground/minced beef (see Jaravata et al.33 as the only published study), which suggests that beef may not be as important a vehicle as cows’ milk. However this study used detection methods developed for analysis of milk rather than beef and a properly validated survey of beef samples is still required to determine if beef is a significant potential vehicle of transmission of Map to humans. Water represents another potential vehicle of transmission of Map to humans. Specifically surface waters have been shown to be contaminated with Map by run-off from pastures grazed by animals with Johne’s disease. Map has been detected by PCR and/or culture in untreated surface water entering water treatment plants in two separate studies in Northern Ireland34 and South Wales, UK.35 Laboratory studies indicate that chlorination has a limited effect on inactivation of Map.3 Chlorination on its own is generally only applied to pristine upland waters and ground waters but is more commonly applied in combination with other water treatment processes for waters likely to have high microbial

loads, hence there is a need to investigate the ability of other water treatment processes to remove or inactivate Map to determine whether contaminated ground waters are potential sources of human transmission. The fact that Map is known to be responsible for an animal disease that results in significant economic losses to agriculture, and there is a question mark remaining over its potential role in human disease, means that there is a need to be able to diagnose infected animals shedding Map in either milk or feces so that the cycle of infection and contamination of both the environment and food can be broken. There is increasing interest amongst milk processors, particularly baby milk manufacturers, in being able to identify Map-infected milk supplies or to monitor the Map status of milk from their suppliers. Ideally a rapid, sensitive detection method for Map would permit producers to quickly assess presence/absence of viable Map in milk. However in the absence of such a commercially available test, PCR currently offers the best alternative for rapid indication of presence/absence of Map but this of course does not indicate the viability of the cells detected. In addition some debate currently surrounds the importance of positive Map PCR results in fecal samples as it is argued that cells may be only transiently present in the gut rather than being established as a pathogen. Given the limits of current testing methods, the fact that results of studies using PCR and/or culture to detect Map in Crohn’s disease patients have varied is perhaps not surprising. A review of the current evidence for and against the relationship between Map and Crohn’s disease can be found in Behr and Kapur.19

17.1.3 Current Methods for the Detection of Foodborne Map 17.1.3.1  Phenotypic Techniques Culture is the “gold standard” method of detecting viable Map in clinical or food specimens. However, Map is the slowest growing of the cultivable mycobacteria, with a generation time under optimal growth conditions of over 20 h,36 so primary culture from veterinary, clinical or food specimens can take 6 months or longer from testing to confirmation of the identity of an isolate as Map. However, once established, a Map isolate will produce colonies within 4–6 weeks upon subculture at the optimum growth temperature of 37oC under aerobic conditions. Colonies are small (1–2 mm), usually white and domed with an entire margin on Herrold’s Egg Yolk medium plus mycobactin J (HEYM); rough and pigmented colonies are rarely seen. Map’s extremely slow growth is generally attributed to its inability to produce mycobactin, a siderophore that is responsible for the binding or transport of iron into cells. Consequently mycobactin J (commercially available from Synbiotics Europe SAS, Lyon, France or Allied Monitor, Inc., MO) must be included in any media used to culture this bacterium in the laboratory. Automated or semi-automated broth culture systems— BACTEC 12B and MGIT (Becton Dickinson) and Trek ESP (Trek Diagnostic Systems) —have now been widely

232

adopted for culture of Map from diagnostic specimens (principally feces) and for milk testing. These offer a more rapid indication of growth of Map (cultures are incubated for up to 40 days) than culture on solid HEYM, but the main deficiency of all these broth culture systems is that frequently mixed cultures of acid-fast and non-acid-fast bacteria and/ or fungal contaminants are obtained. Hence the detection of the growth of Map must be confirmed from culture-positive samples either by sub-culture on to solid agar medium followed by PCR confirmation of any suspect acid-fast colonies that grow, or by PCR testing of a portion of the broth culture following DNA extraction.37,38 Whilst culture remains the only means available to demonstrate the presence of viable Map in a sample the method is problematic for a number of reasons. Firstly, chemical decontamination of the sample is required, the purpose of which is to inactivate the non-acid-fast bacteria present in the sample that would overgrow the mycobacteria during the long incubation period. Commonly used decontaminants include 0.75% cetylpyridinium chloride (CPC, also referred to as HPC) for milk,39,40 and CPC in conjunction with double incubation in Brain Heart Infusion broth and an antibiotic cocktail for feces (Cornell method37). These treatments were developed to not affect the viability of acid-fast cells present in samples, which have more chemically resistant cell walls than other common eubacterial forms. In reality the decontamination step is known to adversely affect Map viability to some degree, resulting in the recovery of only a fraction of the Map cells originally present in a sample leading to an underestimate of either the number of Map present or the number of Map-positive samples. A second problem with this methodology is that if decontamination has not been effective cultures can be overgrown by non-acid-fast bacteria since there is no selective growth medium for Map that would suppress the growth of these organisms. Certain antibiotics can be added to HEYM (vancomycin, amphotericin B, and nalidixic acid) or to broth cultures used in the automated detection systems BACTEC and MGIT (PANTA; polymyxin B, amphotericin B, nalidixic acid, trimethoprim, and azlocillin) to suppress the growth of competitive microorganisms, however this may in turn prevent recovery of Map cells (see Grant et al.41), especially if they are of sub-lethally injured or stressed. Thirdly, the long incubation periods required due to the slow growth of Map necessitates the use of sealed glass bottles instead of Petri dishes, making accurate enumeration of colonies difficult. In summary, culture of Map is deficient in terms of specificity, sensitivity, and speed, the very properties provided by PCR methods for the detection of Map. 17.1.3.2 Bacteriophage-based Techniques Bacteriophage-based rapid methods of identification for M. tuberculosis have been developed42–44 and produced as a commercial assay for the detection of M.tb in human sputum samples (FASTPlaqueTBTM, Biotec Laboratories, Ipswich, UK). This assay detects the replication of a Mycobacteriumspecific phage (D29) rather than the growth of the M.tb cell

Molecular Detection of Foodborne Pathogens

and results are available within 48 h (see Rees and Loessner45). It has recently been demonstrated that the FASTPlaqueTBTM reagents can also be used to detect the presence of viable Map cells in milk.6,46,47 The advantage of using phage to detect the Map cells over conventional culture methods is that the specificity of the phage-host interaction removes the need for chemical decontamination of the milk sample prior to the assay, thus increasing the sensitivity of the test. Some growth of normal milk flora does occur in the agar but this can be reduced by addition of the antibiotic supplement supplied with the FASTplaqueTBTM kit (NOA; nystatin, oxacillin, and aztreonam). To carry out the milk assay 50 ml milk samples are centrifuged to concentrate the mycobacterial cells. These are resuspended in a modified Middlebrook 7H9 medium and phage is added to the sample. After a period of incubation to allow phage infection (1 h) a virucide is added to inactivate any phage that have not infected a mycobacterial cell; the only phage that survive are those that have infected a target cell. The infected cells are then mixed with a lawn of the fast-growing M. smegmatis in a soft agar, and these cells act as hosts to further replicate phage released from the primary infection. After 18 h incubation, plaques are formed in the M. smegmatis lawn indicating the fact that the phage was protected from the virucide and that the sample contained a target mycobacterial cell. As this cell must be viable to allow phage replication the phage test provides live/dead differentiation but does not give specific identification of the Mycobacterium sp. detected. Hence the phage test alone is useful when examining spiked milk samples and can be used to enumerate the number of viable cells present in a sample.6 To allow identification of Map cells in naturally infected milk samples, PCR is used to confirm the identity of the cell detected; DNA is extracted from the plaque and then multicopy signature sequences are amplified by PCR from the single cell detected (in this case IS900; see Section 17.1.3.3 (i)). The combined phage PCR assay sensitively detects viable Map cells in milk samples within 48 h.47 17.1.3.3  Molecular Techniques (i) Single PCR: Once DNA has been successfully isolated from either cultured cells or a processed sample (see Section 17.2.2) the presence of Map can be confirmed by PCR amplification of the Map-specific sequences. PCR methods are currently used to directly detect Map in food samples in food surveillance studies,41 to confirm the presence of Map in broth cultures (Trek ESP II, MGIT, BACTEC) that contain acid-fast bacteria (e.g., Kim et al.,48 Shin et  al.49), to confirm that a suspect acid-fast isolate from solid culture media is Map (e.g., Secott et al.50); and to confirm whether Map or another Mycobacterium sp. has produced a plaque in the FASTPlaqueTBTM phage amplification assay.47 The most commonly used genetic sequence to differentiate Map from other mycobacteria species is the insertion sequence IS900 first reported by Green et al.51. Map has 14–20 copies of IS900, which encodes a 43-kDa DNA binding protein (termed p43). For many years it was thought that IS900 was

Mycobacterium

specific for Map and hence this DNA sequence has been used to design primers for PCR tests and is still routinely used to detect this organism in veterinary, clinical, and food samples or to confirm an acid-fast isolate as Map. However, reports of IS900-like sequences in other mycobacteria began to emerge in the late 1990s and more have been identified as other mycobacterial species have been included in the studies (M. scrofulaceum-type strains,52 M. avium isolates from HIV patients,53 strain 2333 related to M. cookie,54 M. porcinum55) and this has cast doubt on the specificity of IS900 as a signature sequence for Map. IS900 belongs to a family of related insertion elements found in the order Actinomycetales.56 Included in this group are IS901 (specific for M. avium subsp. avium), IS902 (specific for M. avium subsp. silvaticum), IS1110 (found in some M. avium complex isolates) and IS1613 (derived from M. avium subsp. avium) and the greatest degree of homology between these elements is found in the 3′ region of IS900.52 Recently Mobius et al.57 carried out a systematic comparison of different primer sets that have been designed for amplification of IS900 sequences and found that some performed better than others; the most specific single-round PCR assay was found to be the primer sequences published by Bauerfeind et al.58 This was somewhat surprising since the primers were designed to amplify a region in the more conserved 3′ region of IS900 but this group also recommend the use of a high annealing temperature (65°C) to increase stringency and overcome this problem. To avoid issues of primer cross-reactivity, it is common now to use restriction analysis, originally advocated by Cousins et al.52 or DNA sequencing of IS900 PCR products to confirm that the amplicon obtained results from amplification of Map genomic DNA and not from that of another mycobacterial species. It is now widely accepted that IS900 PCR alone cannot be used to test for the presence of Map in food.9 The questionable specificity of IS900 has prompted researchers to seek more specific targets. Genes are highly conserved across the mycobacteria with only a limited number of sequences being identified that can differentiate between Map and other Mycobacterium spp. However several alternatives to IS900 for PCR detection of Map have been reported—F5759, gene encoding the HspX protein60 and ISMav261—see Table 17.1. Recent publication and analysis of the genome sequence of Map has uncovered further new sequences with no homologues amongst other mycobacteria—Locus 251 and Locus 255 (one copy of each, Bannantine et al.62) and ISMap02 (six copies, Stabel, and Bannantine63). The same scale of PCR testing of isolates recovered from food has not been undertaken with these new targets as has been performed using IS900 and therefore their efficacy has not been fully evaluated. However there has been a recent report of non-Map specific sequences being amplified with primers specific for F57 (in M. obuense) and ISMav2 (in M. smegmatis and M. fortuitum).57 (ii) Nested/semi-nested PCR: When using PCR to confirm the identity of cultured Map cells from a suspect acid-fast colony or broth culture, high numbers of Map are likely to be

233

present and therefore sensitivity of PCR assays is not an issue. However when trying to detect low numbers of Map cells from complex samples such as milk, the minimum detection limits of standard PCR methods were considered by some to be leading to false negative results. This is especially a problem when single copy gene sequences are targeted, as the presence of only one copy of the target gene sequence inherently reduces the sensitivity of PCR detection compared to amplification of multicopy insertion sequences such as IS900. To increase detection sensitivity nested or semi-nested IS900 PCR methods were developed.56,64 Nested PCR methods consist of two separate PCR amplifications (30–40 cycles each) involving two different primer sets. The sequence amplified by the second set of primers occurs within the PCR product formed by the first set of primers. In semi-nested PCR one of the primers in the second set is the same as for the first amplification (e.g., primers for F57 PCR described by Vansnick et al.65). Primers for nested PCR methods targeting IS900 are detailed in Table  17.2. These nested IS900 PCR methods have generally achieved a 10–100 fold increase in Map detection sensitivity compared to standard IS900 PCR. However, it must be recognized that the extreme sensitivity of detection when nested PCR is used means that low levels of Map DNA that may be carried over by hands, pipettes, tubes in the laboratory can lead to false-positive results, unless stringent separation and segregation of the different steps in the PCR process and equipment used at each step. A nested/semi-nested PCR approach has also been used when low copy number Map-specific sequences are being targeted (F5765 and ISMap0263), due to the fact that the low concentration of the target sequence inherently reduces the sensitivity of PCR detection compared to the IS900 target which is present in multiple (14–20) copies per Map cell. Published primers for F57 and ISMap02 semi-nested and nested PCR, respectively, are detailed in Table 17.2. Again the specificity and sensitivity of the nested Map-specific PCR detection assays has been recently evaluated by Mobius et al.57 who found that the most sensitive assays had a detection limit of 1 CFU for nested PCR. Interestingly they also reported that the most crucial factor in determining the sensitivity of an assay (single or nested PCR) was the different abilities of the primer pairs to bind to the target DNA under the PCR conditions specified rather than the number of copies of the specific sequence being targeted in the Map genome. This suggests that further optimization of PCR conditions could be carried out for existing primer pairs to increase assay sensitivity. (iii) Real-time PCR: In recent years there has been growing interest in developing real-time PCR methods for detection of Map as it brings the prospect of more rapid and automated detection systems that can be used for routine analysis of samples. Real-time systems use either DNA binding dyes (e.g., SYBR Green) or fluorogenic probes as detailed in Table  17.3. The advantage of fluorogenic probes over DNA binding dyes is that specific hybridization between probe and target is required to generate fluorescent signal, and nonspecific amplification due to mis-priming or primer-dimer

234

Molecular Detection of Foodborne Pathogens

Table 17.2 Published Primer Sets Targeting the Different Map-Specific Sequences used for Conventional and Nested/ Semi-Nested PCR Detection Methods Target IS900 single

IS900 nested

Primer Details (5’ – 3’)

Product Size (bp)

Comments on use, Specificity

Reference

IS900/15OC: CGC TAA TTG AGA GAT GCG ATT GG IS900/921: ATC AAC TCC AGC AGC GCG GCC TCG

229

Amplify IS900-like sequences—strain 2333 related to M. cookii54 M. fortuitum and M. intracellulare, Mobius et al.57

106

P11: CGT CGT TAA TAA CCA TGC AG P36: GGC CGT CGC TTA GGC TTC GA

278

Amplify IS900-like sequences—strain 2333, Englund et al.54; M. scrofulaceum-type, Cousins et al.52

107

P90: TCG GGG CCG TCG CTT AGG P91: GAG GTC GAT CGC CCA CGT GA

400

Amplify IS900-like sequences—M. avium subsp. avium HIV isolates, Naser et al.53; strain 2333, Englund et al.54

108

MK5: TTC TTG AAG GGT GTT CGG GGC C MK6: GCG ATG ATC GCA GCG TCT TTG G

558

Specific for Map but by-products observed (Mobius et al.57)

109

P90+: GAA GGG TGT TCG GGG CCG TCG CTT AGG P91+: GGC GTT GAG GTC GAT CGC CCA CGT GAC

413

Amplify IS900-like sequences—M. scrofulaceum type, Cousins et al.52; M. porcinum, Taddei et al.55

93

MP3c: CTG GCT ACC AAA CTC CCG A MP4c: GAA CTC AGC GCC CAG GAT

314

Amplify IS900-like sequences from strain 2333

58

S204: TGA TCT GGA CAA TGA CGG TTA CGG A S749: CGC GGC ACG GCT CTT GTT

563

Amplify IS900-like sequences from strain 2333

64

1st round: S204/S749 (as above) 2nd round: S347: GCC GCG CTG CTG GAG TTG A S535: AGC GTC TTT GGC GTC GGT CTT G 1st round: TJ1: GCT GAT CGC CTT GCT CAT TJ2: CGG GAG TTT GGT AGC CAG TA 2nd round: TJ3: CAG CGG CTG CTT TAT ATT CC TJ4: GGC ACG GCT CTT GTT GTA GT

64

210

356

Described as “uniquely specific for IS900” and designed to avoid amplification of IS900-like sequences from 2333, WA isolates, etc.

56

294

hspX

Left: GAC CGG CTA TCT GTG GAA C Right: CTC GTC GGC TTG CAC CTG

211

110

F57

f57p1: TTG GAC GAT CCG AAT ATG T f57p2: AGT GGG AGG CGT ACC A

254

77

F57: CCT GTC TAA TTC GAT CAC GGA CTA GA R57: TCA GCT ATT GGT GTA CCG AAT GT F57Rn: TGG TGT ACC GAA TGT TGT TGT CAC R57: as 1st round

432

III20: GAT CAT TCC CGG ATG TGT G III21: AGA CTG CGG TGA AAC TGC T

269

ISMav1: GTA TCA GGC CGT GAT GGC GG ISMav2: CGC GAC CAG CGC TCG ATA CA

318

ISMav2-F: GTG AGT TGT CCG CAT CAG AT ISMavB2: GCA TCA AAG AGC ACC TCG AC

494

49

Locus 251

251F: CAC GTG CTG TCC CCA TCG GC 251R: CTA CGT CTT CGT GAC CAA AG

417

62

Locus 255

255F: CAG TCA CCC CGC GGC CGG TA 255R: TCT ACT GAC CCG CAG ATC GAA

402

ISMav2

False-positive due to M. obuense, Mobius et al.57

65

424 61 Nonspecific products due to M. fortuitum and M. smegmatis reported by Mobius et al.57

By-products reported by Mobius et al.57 with “other mycobacteria”

102

62

(Continued)

235

Mycobacterium

Table 17.2  (Continued) Target ISMap02

Primer Details (5’ – 3’)

Product Size (bp)

Forward: GCA CGG TTT TTC GGA TAA CGA G Reverse: TCA ACT GCG TCA CGG TGT CCT G

278

Forward 2nd: GAT AAC GAG ACC GTG GAT GC Reverse 2nd: AAC CGA CGC CGC CAA TAC G

117

artifacts do not generate signal. Quite a number of different primer sets have been published targeting the range of Map-specific elements and genes (Table 17.3). The ability to monitor the real-time progress of a PCR is a significant advantage. Reactions are characterized by the point in time during cycling (threshold cycle, CT) when amplification of a PCR product is first detected rather than the amount of PCR product at the end of a fixed number of cycles. Real-time PCR can become quantitative if appropriate standard curves are constructed relating CT value to copy number of the target element/gene. Real-time PCR assays are reported to: have equivalent or greater detection sensitivity compared to conventional PCR depending on Map target employed;63,66,69 achieve equivalent detection sensitivity for Map as culture;68,69 be faster than conventional PCR—30–40 mins for 40 cycles compared to a 1–2 h;70,71 and be able to detect multiple targets simultaneously (e.g., IS900 + Locus 251, IS900 + F5772), an approach advocated by Herthnek and Bolske73 to circumvent IS900 specificity issues. (iv) Inclusion of internal Controls in PCR reactions: Increasingly, an internal amplification control (IAC) molecule is being included in PCR reactions so that PCR inhibition can be distinguished from lack of amplification of a PCR product and to allow test results to be validated.74 Contaminants which lead to nonamplification of the organism-specific target result in a false-negative result being recorded. This is very important when processing with samples that contain organic matter, be it milk or feces.75 If the IAC molecule does not amplify and produce the expected PCR product then this is indicative of the presence of inhibitory substances in the sample. Inhibition may be total, resulting in complete reaction failure, or partial, resulting in reduced sensitivity of detection. IAC molecules added to Map PCR reactions have included the human β-actin gene,54,64 the Listeria monocytogenes hly gene,76 the pUC19 plasmid DNA,77,78 and by genetically modifying a plasmid containing an insert of M. bovis-hspX.79 As expected commercially-available PCR detection kits for Map, such as ADIAVET ParaTub PCR (ADIAGENE, St Brieuc, France), BACTOTYPE  Detection kit M. paratuberculosis Milk Test (Labor Diagnostik, Leipzig, Germany), and M. paratuberculosis 209/700IC kit (Sacace Technologies Srl, Caserta, Italy), all incorporate IAC as part of their QC procedures. (v) NASBA: Nucleic acid sequence-based amplification (NASBA) normally involves amplification of an RNA template using temperatures within a narrow range (isothermal;

Comments on Use, Specificity

Reference 63

generally 37–42°C). The reaction uses the enzyme reverse transcriptase to synthesize cDNA from a RNA target sequence, RNase H to digest the RNA strand and then synthesis of up to 100 RNA copies from the cDNA by the enzyme T7 RNA polymerase from the cDNA template. This RNA then acts as a template in the next round of amplification, leading to a rapid increase in the concentration of the amplified product. NASBA requires two primers, one that is homologous to the 3′ end of the target sequence and includes a bacteriophage T7 promoters sequence at the 3′ end to allow second round synthesis of RNA, and a second primer homologous to the 5′ end of the sequence. A NASBA detection assay has been developed that targets the dnaA gene of Map.80 This assay incorporated a molecular beacon probe so that amplification of the target gene could be detected by real-time PCR; this probe is homologous to a sequence located between the two amplification primers and includes a stem-and-loop-structure which is double-labeled with a fluorescent dye at the 5′ end and a universal quencher at the 3′ end. When this probe hybridizes to the target RNA, the fluorescent reporter dye and the quencher are linearly separated and results in an increase in fluorescence intensity proportional to the concentration of the target RNA molecular to which it hybridizes. The fluorescent signal is then quantified using standard real-time PCR methodologies. As molecular beacon probes can be labeled with different fluorescent dyes, a multiplex PCR was developed to allow the simultaneous amplification and detection of the target gene and an IAC molecule.76 The NASBA assay had a sensitivity of approximately 200 cells per reaction, and was able to detect 103 Map cells in artificially contaminated drinking water or 104 Map cells added to pasteurized semi-skimmed milk indicating, again, that contamination by organic matter from the sample matrix is a consistent problem for molecular methods of detection. (vi) Microarrays: The advent of whole genome sequence analysis has led to the development of microarray technology where every gene or every sequence of a bacterial genome is displayed on a solid support in an ordered grid which can then be hybridized with DNA from other species or isolates to allow comparison of whole genome content. This technology is not used for routine identification of organisms but has led to rapid advances in our understanding of the stability of genomes and identification of variable regions that may otherwise have been missed. For instance, genome comparison using microarrays has identified large sequence polymorphisms (LSPs) that are variably present in different strains of Map, and that these

236

Molecular Detection of Foodborne Pathogens

Table 17.3 Published Primer and Probe Sets Targeting Map-Specific Sequences used for Quantitative Real-Time PCR MAP Target IS900

Real-Time PCR Equipment

Primers and Probes (5′–3′)

Reference

Primers of Collins et al. (1993) Probe: TET-CGG ACC GTA ACT ACC CGC GGC GTG ATG GGT CCG-dabcyl

ABI PRISM 7700 Sequence Detection System

70

F2: AAT GAC GGT TAC GGA GGT GGT R2: GCA GTA ATG GTC GGC CTT ACC Probe P2: FAM-TCC ACG CCC GCC CAG ACA GG-TAMRA

ABI PRISM 7700 Sequence Detection System

105

P90: GTT CGG GGC CGT CGC TTA GG P91: GAG GTC GAT CGC CCA CGT GA SYBR green

LightCycler

71

F1: CGG GTA TGG CTT TCA TGT GGT R1: GTC GAT CGC CCA CGT GAC FL-labeled probe: CCA CCT CCG TAA CCG TCA TTG TCC AGA LC Red 640-labeled probe: CAA CCC AGC AGA CGA CCA CGC

LightCycler

111

Forward: CGG GCG GCC AAT CTC Reverse: CAG GGA CGT CGG GTA TG Probe: FAM – TTC GGC CAT CCA ACA CAG CAA CC- TAMRA

GeneAmp 5700 Sequence Detection System

66

IS900QF: CCG GTA AGG CCG ACC ATT A IS900QR: ACC CGC TGC GAG AGC A Probe: FAM-CAT GGT TAT TAA CGA CGA CGC GCA GC-TAMRA

ABI PRISM 7700 Sequence Detection System

112

SF187: TGG TCG TCT GCT GGG TTG A SR239: GCC ACA ACC ACC TCC GTA AC SYBR green

ABI 5700 Sequence Detection System

113

SF214: ATG ACG GTT ACG GAG GTG GTT SR289: TGC AGT AAT GGT CGG CCT TAC Probe PR265: FAM-CGA CCA CGC CCG CCC AGA-TAMRA F57

MAPf57p1: TTG GAC GAT CCG AAT ATG T MAPf57p2: AGT GGG AGG CGT ACC A Probes: MAPf57-3iFluo: CAC GCA GGC ATT CCA AGT MAPf57-5iRed705: TGA CCA CCC TTC CCG TCG (IC probe)

LightCycler

78

Locus 251

251F: GCA AGA CGT TCA TGG GAA CT 251R: GCG TAA CTC AGC GAA CAA CA Probe: FAM-CTGACTTCACGATGCGGTTCTTC-BHQ

SmartCycler II

72

ISMap02

Forward: GCA CGG TTT TTC GGA TAA CGA G Reverse: TCA ACT GCG TCA CGG TGT CCT G

Unclear—MJ Research?

63

Forward 2nd: GAT AAC GAG ACC GTG GAT GC Reverse 2nd: AAC CGA CGC CGC CAA TAC G SYBR green ISMav2

ISMav2 F: GAT GAG TGG GTC GAG GAC TAC AA ISMav2 R: CCG TTG AGC CGG TGT GAT Probe: FAM – CCA AGC CCT AAA GAT - MGB

ABI 7000 Sequence Detection System

114

hspX

Forward hspX Primer: GAC CGG CTA TCT GTG GAA C Reverse hspX Primer: CTC GTC GGC TTG CAC CTG hspX Upstream Probe: GCA CCC GTC GTG GTA TCT-FLU hspX Downstream Probe: LC RED 640-AAT CTG CAA GCC AAT CCG G-PHOSPHORYLATION

LightCycler

79

Key for probe labels: FAM=5-carboxyfluorescein, MGB=minor groove binder,TAMRA=6-carboxy-tetramethyl-rhodamine, FL=fluorescein,TET=tetrachloro6-carboxyfluorescein, dabcyl=4-dimethylaminophenylazobenzoic acid, FLU=fluorescein, BHQ=Black Hole Quencher 1.

LSPs can be the result of either horizontal gene transfer or insertion or deletion events.81 When the Map genome content was compared with other members of the M. avium complex, it was noted that genes predicted to be involved in metabolic

pathways were overrepresented in the LSPs, and that a polymorphism within the mycobactin biosynthesis operon potentially explains the mycobactin J dependency of Map when grown in vitro.82 Similarly Marsh et al.83 used microarrays to compare

237

Mycobacterium

the genomic content of sheep (S) and cattle (C) strains that are known to have different host tropisms. They identified three large genomic deletions in the sheep strain, totaling 29,208 bp and involving 24 open reading frames providing a genetic basis for the different behavior of these host-adapted strains that had been previously reported. These studies have been reviewed, along with other methods of whole genome analysis that are outside the scope of this chapter, by Turenne et al.84

17.2  Methods 17.2.1 Reagents and Equipment Reagents

Consumables

Equipment

Map-specific primers—see Tables 17.2 and 17.3 Lysis buffer

Pipette tips

Centrifuges—50 ml and 1.5 ml capacity

Centrifuge tubes: 50 ml, 1.5 ml

Tris-EDTA buffer

dNTPs

Lysing matrix blue (QBiogene) or Zirconium beads 0.1 mm Sterile pastettes

Bead-beater— FastPrep (QBiogene), BioSpec mini beadbeater, or similar Conventional thermal cycler or real-time PCR system

Taq polymerase

PCR tubes

Phenol:chloroform:isoamylalcohol (25:24:1) Isopropanol 70% alcohol Phosphate-buffered saline containing 0.05% Tween 20 (wash buffer if using IMS) Antibody or peptide coated magnetic beads for IMS

Gel electrophoresis equipment Magnetic rack if using IMS Pipettes

17.2.2 Sample Preparation Sample preparation steps employed for foods are key to providing high purity DNA free from inhibitors and in maximal quantities for PCR purposes. This means concentrating low numbers of Map in milk by centrifugation, removing milk inhibitors by immunomagnetic separation (IMS), peptidemediated capture or magnetic capture and washing, lysing the tough Map cell wall by bead-beating, and purifying and precipitating the DNA released. Due to the low levels of Map expected to be present in milk samples it is usual to test samples of up to 50 ml milk and use centrifugation to concentrate this into a more appropriate sample volume (1 ml) before further processing. The vast majority of published milk surveillance studies have tested pellet only by both culture and PCR.26,41,66,85–92

However, there is some debate about how Map segregates into the three milk fractions (cream, whey, and pellet) upon centrifugation and, therefore, whether testing the pellet ensures adequate detection sensitivity or whether cream and pellet should be combined before further processing. Millar et al.93 were the first to comment on segregation of Map in milk fractions following centrifugation. These researchers found that disaggregated intact Map spiked into whole pasteurized cows’ milk gave an IS900 PCR signal which segregated into both the cream and pellet fractions after centrifugation, although a much stronger PCR signal was obtained from the pellet fraction, whereas when milk was spiked with Map DNA in the same study a PCR signal was observed equally in all three milk fractions. When Millar et al.93 applied IS900 PCR to naturally infected raw milk samples IS900 PCR signals were observed in all three milk fractions, but most consistently the pellet and cream fractions tested positive for Map. Gao et al.40 also reported that more live Map could be recovered from the pellet of heat treated milk than from raw milk, although they also state that spiked Map cells preferentially partitioned into cream in raw milk. In a subsequent study Gao et al.94 advised heat treatment of 50 ml raw milk samples in a 95oC waterbath for 10 min until the internal temperature reaches over 85oC followed by cooling in ice water for 10 min prior to DNA extraction to maximize sedimentation of Map in the pellet fraction, resulting in stronger PCR signal and less background signal (smearing). They also reported that if the cream and pellet fractions obtained from raw milk were treated with 0.75% hexadecylpyridinium chloride (the usual decontaminant prior to culture) for 30 min at room temperature before further centrifugation and discard of the supernatant and cream layer then the PCR signal in the resultant pellet increased. Pooling of pellet and cream fractions prior to further extraction definitely improved PCR detection sensitivity according to Gao et al.94 as did inclusion of 0.0037% bovine serum albumen in the PCR reaction.95 C-18-carboxypropylbetaine (CB-18) has been tried as an aid to sedimentation of Map from milk96 but no improvement over a dilution and centrifugation method was observed for Map, whereas CB-18 had been shown to aid sedimentation of M. bovis from milk previously.97 Once Map cells have been concentrated by centrifugation, the pellet, or pellet and cream, can either be lysed directly or subjected to IMS, ­peptide-mediated capture or magnetic bead capture to selectively separate Map from milk components that may prove inhibitory to subsequent PCR before lysis. The first IMS for Map was developed by Grant et al.98 using rabbit anti-Map polyclonal IgG antibody secondary coated on to Sheep anti-rabbit IgG Dynabeads (Dynal). IMS achieves two things; the simultaneous selection of Map from a mixed background microflora and removal of milk components (such as Ca2+ ions, proteins, thermonucleases75) that may inhibit PCR. The use of the original IMS method in conjunction with IS900 PCR (IMS-PCR) was subsequently described by Grant et al.99 for Map in milk and further improvements to this method (inclusion of bead beating and DNA purification steps) to improve PCR detection sensitivity were described by Khare et al.66 An IMS method employing monoclonal antibody capture has also been described100 and is now part

238

of a commercial IMS-real-time PCR kit for Map in milk (AnDiaTec ParaTub-SL IMS-Taqman Real-time PCR kit, AnDiaTec GmbH & Co KG, Korwesheim, Germany). Stratmann et al.101,102 used Map-specific peptides aMP3 and aMptD rather than antibody to achieve magnetic capture, with similar or greater detection sensitivity by PCR than antibody capture, and peptide capture is now part of another commercial detection kit for Map in milk (BACTOTYPE M. paratuberulosis milk test, Labor Diagnostik, Leipzig, Germany). In contrast, the commercially available Adiapure DNA extraction kit (Adiagene) incorporates magnetic capture with uncoated beads (identity unknown), which have been found to work just as well for concentrating Map from milk samples as the original polyclonal antibody-based IMS method in a recent study.67 Probably the hydrophobic nature of Map (and other mycobacterial) cells make them physically attracted to beads of a certain composition. We have certainly observed significant levels of nonspecific binding of several Mycobacterium spp. to different types of uncoated magnetic beads, including Pierce MagnaBind carboxyl derivatized beads (Grant et al., unpublished data,). Once concentrated from milk (by centrifugation alone or by magnetic separation) Map cells must be lysed to release DNA for PCR purposes. The cell wall of Map has been described by one researcher (Prof. John Hermon-Taylor) as “teflon coated” and boiling alone (95–100oC for 15 min), a method that proves adequate for the lysis of many other bacteria, is generally not sufficient for Map lysis unless large numbers of Map are present in a sample (e.g., when confirming acid-fast cells present in a broth culture or from a colony to be Map by PCR). However, in situations where low numbers of Map are suspected/expected boiling is not sufficient for lysis. A variety of approaches have been studied, including the use of commercial DNA extraction kits (e.g., InstaGene Matrix, QIAamp Tissue kit, Adiapure) but the consensus opinion is that bead-beating in some shape or form is necessary for maximal release of Map DNA. The two main disruptor systems used for Map are the Bio101 FastPrep Cell Disrupter instrument (Qbiogene), also known as the Hybaid Ribolyser, and the Mini Bead-beater (BioSpec). Bluecapped Lysing matrix B tubes (QBiogene) are used with the FastPrep machine and 0.1 mm zirconia/silica beads (BioSpec) are used with the mini beadbeater. Bead-beating is generally preceded by resuspension of the pellet or magnetic beads with cells attached in a lysis buffer, typically containing Tris, EDTA sodium chloride and sodium dodecyl sulphate at pH 8, and treatment with proteinase K to weaken the cell wall, followed by isopropanol precipitation of DNA. Odumeru et al.94 compared a number of extraction protocols and reported a PCR detection limit of 105 CFU/ml with boiling alone which increased to 102 CFU/ml if cells were subjected to bead-beating (BioSpec mini bead-beater), boiling and isopropanol precipitation instead. Sequence capture following lysis to pull out Map-specific sequences using biotin-labeled specific probes and streptavidin-labeled Dynabeads has also been investigated.103,104 Whilst results demonstrate that sequence capture works for Map—Vansnick et al.104 report that sequence capture PCR has the same sensitivity as fecal culture for detection of Map in feces (100 Map/g feces)—this approach has not

Molecular Detection of Foodborne Pathogens

been adopted for milk in the same way as magnetic separation prior to lysis to date. In summary, a combination of, at least, centrifugation, harsh lysis, physical grinding (bead-beating), and nucleic acid purification are needed to facilitate maximal extraction of Map DNA from milk and other food matrices. Additional steps might include magnetic separation or sequence capture to more specifically extract Map DNA only.

17.2.3  Detection Procedures (i) Typical DNA extraction protocol: (1) Centrifuge 50 ml of milk at 2,500 × g for 15 min. Discard whey fraction and resuspend cream and pellet in 1 ml PBS containing 0.05% Tween 20 (PBS-T; Sigma P3563). Transfer samples to 1.5 ml microcentrifuge tubes using a pastette. (2) If applying immunomagnetic dispense an appropriate volume of beads (10 µl) into each sample. Vortex each tube briefly then mix gently for 30 min at room temperature on a Dynal mixer or flat-bed mixer. (3) Transfer tubes to magnetic rack for 10 min in order to capture beads then carefully pipette off the fluid leaving the beads adhering to the back of the tube. (4) Wash beads twice with 1 ml PBS-T with capture for 2 min on magnetic rack between washes. (5) Resuspend immunomagnetic beads (IMB) after immunomagnetic separation in 700 µl TEN lysis buffer (2 mM EDTA, 400 mM NaCl, 10 mM TrisHCl, pH 8.0, 0.6 % SDS) containing 20 µg proteinase K (Sigma) and incubate overnight at 37°C. (6) After overnight proteinase K treatment, transfer samples to FastPrep Lysing Matrix B (blue-capped) tubes (QBiogene). (7) Place tubes in the Bio101 FastPrep cell disruptor and mechanically disrupt cells by vibration for 45 sec at 6.5 m/sec, then remove tubes and place on ice for 15 min to allow foam to settle. (8) Add 700 µl phenol:chloroform:isoamylalcohol (25:24:1) pH 8.0 (Sigma) to each blue-capped tube and vortex for 1 min. (9) Centrifuge samples at 10,000 rpm in a benchtop microfuge for 10 min and then transfer top aqueous layer to a fresh 1.5 ml microcentrifuge tube containing 400 µl (0.6 vol) isopropanol (Sigma) using a Pasteur pipette to precipitate DNA (NB. IMB will sediment at bottom of phenol layer in the process). Place tubes in –20°C freezer for at least 30 min. (10) Recover DNA by centrifuging at 10,000 rpm in a benchtop microfuge for 10 min. Wash DNA pellet once with 500 µl 70% ethanol centrifuging in a benchtop microfuge at 10,000 rpm for 10 min. (11) Decant ethanol carefully and allow DNA pellet to air dry briefly before resuspending DNA in 50 µl Tris-EDTA (TE) buffer pH 8.0. (12) Perform IS900 PCR using 5 µl of template DNA per reaction.

239

Mycobacterium

1

2

3

- 1000 bp

- 500 bp

- 100 bp

Figure 17.1  PCR amplification of IS900 element. Agarose gel (1.5% v/w) electrophoresis of IS900 PCR amplicon (400 bp) amplified from purified Map ATCC 19851 DNA using P90/P91 primers and protocol described in Section 17.2.3 (ii) (lane 1). Template DNA was prepared by resuspending one colony of Map in 1 ml distilled water in a sterile 1.5 ml Eppendorf tube and boiling for 20 min. Samples were then centrifuged at 13,000 × g for 3 min, and the supernatant extracted and frozen to –20ºC as a positive control DNA sample). As a negative control, template DNA was prepared in the same way from M. smegmatis cells (lane 2). The band at the bottom of this sample running at ~100 bp represents amplification of nontemplate specific primer dimer; this will also appear in positive samples but is most intense in absence of specific template for primers used in PCR reaction. Molecular weight markers (lane 3) are Promega 100 bp ladder (PR-G2101).

(ii) IS900 Single PCR procedure: In Section 17.2.1 (i) the different PCR targets and primer pairs that have been characterized to date are reviewed, including a critique of their relative specificity. Despite some criticisms, IS900 PCR is still the most commonly performed diagnostic PCR assay performed and a suggested protocol is included below. Readers should be aware that buffer composition, thermal cycle programmes, and expected product sizes all differ depending on the primer set chosen; see Table 17.2 and the individual publications cited for PCR protocols for alternative targets/primer pairs. Suggested PCR protocol: (1) Prepare sufficient PCR supermix for the number of PCR reactions required (i.e., no. of samples+positive, control+negative control) using the following components per 45 µl reaction:

Molecular grade water 0.037 % (w/v) Bovine serum albumin

26 µl 5 µl

10x Taq buffer without MgCl2 MgCl2 (50 mM) Primer P90* (30 pmol/µl)

5 µl 1.75 µl 2 µl

Primer P91* (30 pmol/µl)

2 µl

dNTP mix (25 mM of each NTP) Platinum Taq DNA polymerase** (5 U/µl)

3 µl 0.25 µl

* Primers of Vary et al.106. Sequences detailed in Table 17.2. ** Hot Start Taq DNA polymerase available from Invitrogen.

(2) Dispense the supermix in 45 µl aliquots in 0.5 ml PCR thermotubes and add 5 µl sample DNA template, 5 µl molecular biology grade water (negative control) or 5 µl of DNA from a confirmed culture of Map lysed by boiling (positive control). (3) Place tubes on thermal cycler and run the following program: one cycle of 94oC for 2 min, 62oC for 15 sec, and 72oC for 1 min; 35 cycles of 94oC for 30 sec, 62oC for 15 sec, and 72oC for 1 min; and one cycle of 94oC for 30 sec, 62oC for 15 sec, and 72oC for 5 min. (4) Visualize PCR product by 1–2 % (w/v) agaraose gel electrophoresis using 15 µl PCR product mixed with 5 µl loading dye per lane. Run gel at 100 V for 1–1.5h. (5) Confirm the presence of expected Map-specific 400 bp IS900 product by staining gel with ethidium bromide and viewing on UV transilluminator (see Figure 17.1).

17.3 Conclusions and Future Perspectives Molecular detection methods for Map have come a long way in recent years as there has been intense research effort to improve and optimize PCR detection methods for Map in milk. This research has been driven by the potential link between Map and Crohn’s disease and the potential need to control Johne’s disease in the food producing animal population and/or monitor Map contamination of animalderived foods by food producers and processors. Current PCR methods permit the rapid, sensitive, and specific detection of Map DNA in food although the origin of this DNA—whether from live or dead Map—is still not differentiable by PCR methods. Accurate quantitation of viable Map levels existing in naturally infected raw milk would assist risk assessment and enable prediction of process efficiency (pasteurization) in the food industry. Whilst real-time PCR methods now afford the possibility to quantify Map in a milk sample, Map viability is not confirmed by real-time PCR unless the concentration of PCR product is shown to increase over time, i.e., minimum two PCR reactions at different time points, during incubation.105 Efforts to detect and quantify RNA as opposed to DNA of Map have to date not been very successful. Our recent work describing a phagebased test to detect viable Map cells in milk demonstrates that rapid, sensitive, and specific detection of Map cells is

240

possible and that further development of these assays may produce practical tests that can be used both for veterinary diagnosis of disease and screening of suspected food vehicles of transmission.

References













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18 Staphylococcus

Paolo Moroni, Giuliano Pisoni University of Milan

Paola Cremonesi, Bianca Castiglioni

Institute of Agricultural Biology and Biotechnology

Contents 18.1 Introduction.................................................................................................................................................................... 245 18.1.1 Morphology and Biology.................................................................................................................................. 245 18.1.2 Pathogenesis and Clinical Features.................................................................................................................. 246 18.1.3 Conventional Testing........................................................................................................................................ 247 18.1.4 Molecular Testing............................................................................................................................................. 247 18.1.4.1 Standard PCR.................................................................................................................................. 247 18.1.4.2 Multiplex PCR ................................................................................................................................ 249 18.1.4.3 Real-Time PCR ............................................................................................................................... 250 18.2 Methods.......................................................................................................................................................................... 252 18.2.1 Reagents and Equipment................................................................................................................................... 252 18.2.2 Sample Collection and Preparation.................................................................................................................. 252 18.2.2.1 Sample Collection............................................................................................................................ 252 18.2.2.2 Sample Pretreatment........................................................................................................................ 253 18.2.2.3 DNA Extraction............................................................................................................................... 253 18.2.3 Detection Procedure......................................................................................................................................... 253 18.3 Conclusions and Further Perspectives............................................................................................................................ 254 References.................................................................................................................................................................................. 255

18.1  Introduction 18.1.1 Morphology and Biology Being a Gram-positive coccus in the genus Staphylococcus, Staphylococcus aureus tends to be arranged in irregular clusters or grape-like clusters when viewed through a microscope and has large, round, golden-yellow colonies, often with hemolysis, when grown on blood agar plates. The golden appearance is the etymological root of the bacteria’s name: the name derived from Greek staphylo (bunch of grapes) and aureus means “golden” in Latin. S. aureus is a facultative anaerobe (fermentative) with the following main features: (i) it can grow at an aw of 0.86 and pH above 4.81 and within the temperature range 7–48°C with an optimum of 35–40°C; (ii) it is catalase positive (meaning that it can produce the enzyme “catalase”) and able to convert hydrogen peroxide (H2O2) to water and oxygen, which makes the catalase test useful to distinguish staphylococci from enterococci and streptococci; (iii) it is oxidase negative; and (iv) it is nonmotile. S. aureus can be differentiated easily from most other staphylococci by its ability to produce coagulase and a heat-stable endonuclease. S. aureus is primarily coagulase-positive (CPS) (meaning that it can produce the enzyme “coagulase” that causes clot formation) while most other Staphylococcus

species are coagulase-negative (CNS). However, while the majority of S. aureus strains are CPS, some may be atypical and do not produce coagulase enzyme. Members of the S. aureus species produce a number of extracellular compounds including membrane-damaging toxins, epidermolytic toxin, toxic shock syndrome toxin, pyrogenic exotoxin, exoenzymes (coagulase and thermostable nuclease) and staphylococcal enterotoxins (SE). The natural reservoirs for S. aureus are the skin and the mucous membrane of humans and animal. Many staphylococcal species have become adapted to life on particular animal species but, in contrast, S. aureus is present on most marine and terrestrial mammals, and may be present as a nonaggressive member of the normal skin microflora or may be associated with infectivity and disease. Up to 30–50% of the human population are carriers of S. aureus; the body sites most often colonized are the nostrils, throat, hair, and hands. This organism may also be isolated from healthy domestic and food animals as well as being associated with disease, particularly mastitis. S. aureus strains have been classified into biotypes according to their human or animal origin: six different biotypes (human, non-β-hemolytic human, avian, bovine, ovine, and nonspecific) have been identified based on biochemical characteristics.2 245

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Molecular Detection of Foodborne Pathogens

18.1.2  Pathogenesis and Clinical Features Staphylococcal food poisoning (SFP) relies on one single type of virulence factor: the SEs. The SEs are a group of extracellular protein with molecular weights in the range of 27000–30000 Daltons, and similar composition and biological activity. To date, 19 types of staphylococcal SEs have been described.3 On the basis of their antigenicity and mode of action in the host, SEs have been divided into two groups. The members of group 1, classical emetic toxins designated SEA, SEB, SEC1, SECbov, SED, and SEE, are the cause of about 95% of SFP in humans.4 These toxins are resistant to gastrointestinal proteases and after their intestinal absorption, the patient can develop symptoms of food poisoning, depending on the SE amount ingested.4,5 Group 2, includes toxins possibly involved in the remaining 5% of SFP outbreaks, i.e., the recently identified SEs, designated SEG, SEH, SEI, SEJ, SEK, SEL, SEM, SEN, SEO, SEP, SEQ, SER, and SEU,6–12 the emetic activities of which are not currently fully understood (Table 18.1). The symptoms of SFP are abdominal cramps, nausea, vomiting, sometimes followed by diarrhea.6 The onset of symptoms is rapid (from 30 min to 8 h) and spontaneous remission is usually observed after 24 h. Hospitalization is required in approximately 10% of the cases.13 In all cases of SFP, the foodstuff or one of the ingredients was contaminated with a SE-producing S. aureus strain and was exposed, at least for a while, to temperatures that allow S. aureus growth. Most of the time the foodstuff reaches this temperature because of a failure in the refrigeration process, or because a growth-permissive temperature is required during processing (e.g., cheese making). Many different foods can be a good growth medium for S. aureus, and have been implicated in SFP, including milk and cream, cream-filled pastries, butter, ham, cheeses, sausages, canned meat, salads, cooked meals and sandwich fillings. The foods that are most often involved in SFP differ widely from one country to another. In the United Kingdom,

Table 18.1 Major Characteristics of 18 Staphylococcal Enterotoxins (SE) SE Type A B C1 C2 C3 C (bovine) C (sheep) C (goat) D E G

Reference [119] [120] [121] [122] [123] [124] [124] [124] [125] [126] [127]

SE Type H I J K L M N O P Q R

Reference [128] [127] [129] [130] [131] [7] [7] [7] [132] [133] [134]

U

[135]

for example, 53% of the SFP reported between 1969 and 1990 were due to meat products, meat-based dishes, and especially ham; 22% of the cases were due to poultry, and poultry-based meals, 8% were due to milk products, 7% to fish and shellfish and 3.5% to eggs.14 In France, in a 2-year period (1999–2000), among the cases of SFP in which the food involved had been identified, milk products and especially cheeses were responsible for 32% of the cases, meats for 22%, sausages and pies for 15%, fish and seafood for 11%, eggs and egg products for 11% and poultry for 9.5%.15 In the United States, among the SFP cases reported between 1975 and 1982, 36% were due to red meat, 12.3% to salads, 11.3% to poultry, 5.1% to pastries and only 1.4% to milk products and seafoods. In 17.1% of the cases, the food involved was unknown.16 During a 5-year period (1988–1992), S. aureus caused 5.1% of the food poisoning outbreaks reported in Europe17 and in Italy it caused four of the 233 outbreaks reported. S. aureus has been isolated from several foods: meat and meat products, chicken, milk, and dairy products, fermented food items, vegetables, fish products, etc.14,18 Salted food products, such as ham, have been reported to be responsible for about 24% of all the cases of staphylococcal intoxication.19 Thus, the origins of SFP differ widely among countries; this may be due to differences in the consumption and food habits in each of the countries. In France, for example, the consumption of raw milk cheeses is much higher than in Anglo-Saxon countries. This may explain the relative importance of milk products involved in SFP in France. In any case, the main sources of contamination are humans (handlers contaminate food via manual contact or via the respiratory tract by coughing and sneezing), and contamination occurs after heat treatment of the food. Nevertheless, in foods such as raw meat, sausages, raw milk, and raw milk cheese, contaminations from animal origins are more frequent and due to animal carriage or to infections (e.g., mastitis). Some S. aureus strains, so-called ‘‘endemic strains’’, are present in some processing plants, such as poultry processing lines,20–22 consequently food products may originally become contaminated during or after processing. However, the presence of large numbers of staphylococci is not sufficient cause to incriminate a specific food as the vehicle of food poisoning because not all staphylococci are enterotoxigenic. In addition, demonstration of enteroxigenicity of food isolates is only circumstantial evidence of enterotoxigenic staphylococcal contamination and the potential for causing food poisoning cannot be ascertained without demonstrating the actual presence of the SE in a suspect food. Conversely, neither the absence of S. aureus nor the presence of small numbers of bacteria is complete assurance that the food is safe (because the SE is extremely resistant and may survive processes which kill the bacterial cells). Concerning S. aureus in food, it should be noted that only the enterotoxinogenic strains possess a risk to public health and the criteria applied should prevent the production of SE during processing and the occurrence of SE in the final product. It is generally considered that enterotoxinogenic staphylococci must reach levels of at least 105–106 cfu/g or ml to

Staphylococcus

produce detectable amounts of SE and, in Europe, low degree contaminations by S. aureus are tolerated in most foodstuffs (up to 104 cfu/g in raw milk cheeses, in France), as they are not considered a risk for public health.

18.1.3 Conventional Testing Risk assessment in foodstuffs relies on classical microbial detection and quantification of CPS staphylococci on a selective Baird-Parker medium, whose composition is standardized (EN ISO 6888/1 and 2).23,24 The sensitivity of these routine tests is around 102 cfu/g for solid foodstuffs and 10 cfu/g for liquid samples. The different media used for the detection and quantification of S. aureus have been reviewed by Baird and Lee.25 Food testing using microbiological criteria may have limited usefulness for food safety for a number of reasons, including low prevalence of the pathogen, or low diagnostic sensitivity of the testing procedure applied. While the finding of a pathogen in a foodstuff may indicate a problem for public health, necessitating appropriate risk management action; the failure on the other hand to detect a pathogen in a food product does not necessarily mean that the pathogen is absent from that food product, process or food lot.

18.1.4 Molecular Testing Use of DNA-based assays may circumvent some of the problems associated with conventional microbiological procedures. Perhaps the greatest single advantage of DNAbased diagnostic assays is that these methods focus on the unique nucleic acid composition of the bacterial genome rather than on phenotypic expression of products that nucleic acids encode. Therefore, DNA-based identification assays are subject to less variability compared with diagnostic methods based on phenotypic characterization, allowing reliable detection and quantification down to one single nucleic acid target per PCR sample. Moreover, not only the presence of the pathogen but also of the genes encoding for SEs production is important to evaluate as enterotoxins nonproducing strains may also occur. 18.1.4.1 Standard PCR Many polymerase chain reaction (PCR) protocols have been developed for the direct detection or for identification of S. aureus in milk and dairy products.26–33 These PCR methods allow identification of bacteria within hours. However, the sensitivities of PCR assay often vary with different DNA extraction methods, suggesting that the DNA extraction protocols are important for optimization of the assay if it is to be applied to food.34 Consequently, the development of a sample preparation strategy that can effectively sequester high-quality DNA of the pathogenic bacteria from food samples before PCR amplification is needed. Indeed, molecular procedures require highly purified template DNA.35,36 In addition, the PCR-based detection of pathogens is made more difficult when raw material with high

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level of background microflora or complex food matrices are involved.18,37 First, these difficulties could be due to small concentrations of the pathogenic DNA present in a typical sample. Second, various factors affect DNA recovery, including the degree of cellular lysis, binding of DNA to particulate material, and degradation or shearing of DNA. Furthermore, in the case of Gram-positive bacteria such as S. aureus and streptococci, an optimal sample processing method should efficiently lyse resistant bacterial cell walls without damaging target DNA. In addition, direct detection of pathogenic bacteria in food samples38 is hampered by the presence of PCR-inhibitory substances frequently associated with the food matrix itself.39 Particularly in milk, components such as Ca2 + , proteinase, fats, and milk proteins may block DNA and shield it from access by polymerase.40 Breaking down casein and the casein micelle with the enzyme pronase would allow better access to the bacteria for lysis by lysozyme and proteinase K.35 Moreover, cheeses are often regarded as difficult matrices to be assayed by PCR because of the high fat content18,40 which is reported to be one of the major factors reducing the sensitivity of the PCR assay.39 Finally, it is possible that the poor sensitivity of detection of S. aureus is caused by the abundance of DNA extracted from the background microflora of the raw milk acting as inhibiting competitor in PCR reactions. In this case false positive results may occur because of parallel amplification of target genes, such as the regions coding for ribosomal RNA (rRNA), from closely related species.41 To overcome PCR inhibition problems and to increase the sensitivity of the assay, overnight selective-enrichment was frequently used to increase the level of pathogen detection.31,32,34,38 After the enrichment step, sufficient bacteria were present to allow pathogens to be detected when the original sample had as little as 1 cfu/ml. The higher sensitivity of the assay may be due to the dilution of inhibitory substances in the enrichment broth and the increased number of organisms.32 In milk samples, another effect of enrichment before PCR may be the reduced detection of DNA from nonviable bacteria. However, although the prior enrichment step increases the target concentration, it precludes quantification.36,42,43 The vast number of procedures used for this purpose and articles published focusing on this topic indicate that these problems are still far from being solved.44–49 Therefore, the separation and concentration of foodborne pathogens directly from food samples without culture enrichment represent two of the most important aspects of sample preparation. Methods for separating bacteria directly from a food matrix and then concentrating them depend on several chemical, physical, and biological principles. Numerous methods for isolating bacterial DNA directly from milk have been reported in the literature and involve a wide variety of substances including Chelex-100,28 spin columns,29,32,33 lysozyme and proteinase K,31 diatomaceous earth,50 alkaline extraction,30 and pronase.48,51 Methods that use proprietary reagents such as Insta-Gene Matrix (BioRad) and PrepMan Ultra reagent (Applied Biosystems) were also evaluated. Although these

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reagents were faster and more convenient than other methods evaluated, results were not consistent and reproducible when trying to isolate bacterial DNA from milk.52 The method first described by Allmann and coworkers48 and modified by Hein coworkers51 was one of the most consistent and reproducible system evaluated for isolation of bacteria directly from whole milk samples. Even a buoyant density gradient centrifugation were recently used to separate bacteria from complex food matrices, as well as to remove compounds that inhibit rapid detection methods, such as PCR, and to prevent false-positive results due to DNA originating from dead cells.53 A specific, sensitive, and rapid method to extract DNA directly from the main Gram-positive bacteria (including S. aureus) found in milk and raw cheese samples was also described by Cremonesi and coworkers.54,55 The DNA extraction method is based on the lysing and nuclease-inactivating properties of the chaotropic agent, guanidinium thiocyanate, together with the nucleic acid-binding properties of the silica particles. In absence of PCR inhibitors, the level of sensitivity achieved for S. aureus detection were 10 cfu/ml in milk and 100 cfu/g in cheese. Finally, PCR detection is often hampered by the presence of inflammatory somatic cells. To eliminate this problem, a method that allows the effective separation of bacterial cells from somatic cells in mastitis milk with amino-silica was recently established.56 The authors identified the conditions under which most of the somatic cells were adsorbed and only S. aureus desorbed from amino-silica upon addition of a desorption solution; in this way the procedure effectively eliminated somatic cells in heavily contaminated milk samples, which resulted in improved clarity of the PCR band. The major advantages of PCR lay in the possibility of using only few nanograms of nucleic acid samples, allowing the elimination of culture, rapidity, and easy analysis. Several PCR methods were described for the identification of S. aureus,57,58 and staphylococcal strains harboring copies of enterotoxin genes can be rapidly detected by PCR in food samples.59 In these works, PCR amplification of DNA regions coding for rRNA was often used because of the presence of hypervariable regions, which facilitates the design of highly specific oligonucleotide probes and common regions for the design of universal probes. Moreover, rDNA is present in many copies, which permits signal enhancement.29,60 In addition to the 16S rDNA gene, the well established standard target for the identification of bacterial species,61 the sequence of the 16S–23S rDNA spacer region and the 23S rDNA gene have proven useful for identification of S. aureus at the species level.33,41,62 For example, a 23S rDNA-targeted PCR-based system for detection of S. aureus in meat starter cultures and dairy products was developed.41 More recently, using species-specific primers derived from 16S and 23S rDNA, PCR detection was performed with S. aureus isolates. The detection limit of this assay for milk products was 16 cfu/µl, representing a valid diagnostic tool for the detection of milk pathogens in dairy products.63

Molecular Detection of Foodborne Pathogens

Other PCR assay for the detection and identification of this pathogen was established and validated using three conserved genes as the DNA target of S. aureus: fmhA (coding for a factor of unknown function), catalase and femA (coding for a factor essential for methicillin resistance). All the three assay systems showed a detection limit of 100 cells per 20 ml reaction assay.64 The protocols developed could be used for rapid and specific detection of this pathogen in food and environmental samples, especially milk.64 Following the recent publication of the genome sequences of nine S. aureus strains (http://www.ncbi.nlm.nih.gov/ genomes/lproks.cgi), the vicK gene of S. aureus that relates signal transduction of the pathogen was investigated as a target for rapid detection and identification.65 The PCR assay allowed amplification of a 289 bp DNA fragment only from S. aureus and not from other Staphylococcus species and other common bacteria tested. The identification of enterotoxigenic S. aureus strains is usually based on amplification of the coa gene.66–68 Also, the presence of the thermostable nuclease gene (nuc) showed strong correlation with enterotoxin production and it is a marker of food contamination with enterotoxigenic S. aureus.18,69 Furthermore, Kalorey and coworkers70 characterized genotypically by PCR 37 enterotoxigenic strains of S. aureus isolated from 552 milk samples using not only primers that amplified genes encoding coagulase (coa) or thermonuclease (nuc), but also clumping factor (clfA), enterotoxin A (entA), and the gene segments encoding the immunoglobulin G binding region and the X region of protein A gene (spa). To survey the enterotoxin genotypes for S. aureus strains isolated from food-poisoning cases in Taiwan, PCR primers specific for all SE genes, including SEN, SEO, SEP, SEQ, SER, and SEU, or staphylococcal superantigens genes (SAg), including toxic shock syndrome toxin I (TSST-1) genes, were more recently developed.71 Using these PCR primers the genotypes of 147 S. aureus strains isolated from patients associated with SFP outbreaks occurred during 2001–2003 was assayed. 91.8% of these strains were found positive for one or more SE or SAg genes. To define PCR-based detectability of S. aureus in the early stages of raw milk cheese making, raw milk artificially contaminated by different concentrations of S. aureus FRI 137 strain, harboring nuc, sec, seg, seh, and sei genes was employed in dairy processing resembling traditional raw milk cheese making.36 Samples of milk and curd were PCRanalyzed after DNA extraction by targeting all the above genes. The pathogen was detected when the initial contamination was 104 cfu/ml by amplification of nuc and seh genes. 105 and 107 cfu/ml were needed when seg or sei and sec genes were targeted, respectively. Enrichment cultures from raw milk and curd samples provided to increase the detection limit of 1 log on average. Therefore, the direct detection of the pathogen in the raw material and dairy intermediates of production can provide rapid results and highlight the presence of loads of S. aureus potentially representing the risk of intoxication. However, every target gene to be used in the analysis has to be studied in advance in a system similar to

Staphylococcus

the real case in order to determine the level of contamination that can be predicted. Often also the consumption of ham or meat containing SE is identified as the cause of illness. Evaluation of the occurrence of most known SE (SEA to SEE; SEG, SEH, SEI, SEM, SEJ, SEN, and SEO) genes, egc (enterotoxin gene cluster containing the following sequence of genes: seo, sem, sei, phient1, phient2, sen, and seg) and tsst1 (toxic shock syndrome toxin 1) gene in both CPS and CNS staphylococcal strains isolated from meat and dairy products were carried out by Blaiotta and coworkers.72 PCR detection methods were used to analyzed 109 wild Staphylococcus spp. strains isolated from Napoleitype salami, raw water buffalo milk and natural whey cultures used for mozzarella cheese manufacturing, revealing that the occurrence of SE genes in CNS and other non-S. aureus species in these foodstuffs is very rare. To gain insight into the prevalence of S. aureus and its emetic enterotoxins in raw pork and uncooked smoked ham, samples of raw pork, salted meat and ready-for-sale uncooked smoked ham were examined for the prevalence of S. aureus and SEA–SED.60 To this end classical cultural methods were employed as well as molecular biological techniques (PCR) and the results were compared. Fresh meat was contaminated most often. By PCR, 62.2% were identified as being S. aureus positive compared to 57.7% positive samples using the cultural technique. The detection rate decreased significantly during the fabrication process. The pathogen was cultivated from 8.9% of the salted meat samples. Here, 55.6% of the samples reacted positively in the PCR, and finally, in approximately a third of the ready-for-sale smoked hams, S. aureus genes were found. This study clearly shows that the PCR assay is more sensitive than the classical cultural method. Approximately 35% of the staphylococcal strains identified using the PCR technique were enterotoxigenic. Using the SET-RPLA, a percentage of 28.6% enterotoxigenic isolates was ascertained. The detection of SE-genes by PCR is resulted faster and easier to perform than the SET-RPLA. 18.1.4.2  Multiplex PCR The DNA-based identification systems could be targeted to allow for simultaneous rapid screening of a large number of pathogens. For example, rapid and sensitive detection techniques for foodborne pathogens are important to the food industry. In multiplex PCR, multiple pairs of primers specific for different DNA segments are included in the same reaction to enable amplification of multiple target sequences in one assay. In many cases, more than four pairs of PCR primers can be used.73 Primers used in multiplex PCR amplification are chosen to have similar melting temperatures (Tm) as a difference of more than 10°C in the Tm of the two sets of primers may result in differential yields of amplification products,74 and no visible amplification for one or the other target. The major advantage of multiplex PCR over conventional PCR is its cost effectiveness. It reduces the amount of reagents, such as Taq DNA polymerase, used for each diagnosis. Moreover, it requires less preparation and analysis

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time than systems in which several tubes of simplex PCR are used. Phuektes and coworkers32,33 developed a multiplex PCR for detection of S. aureus, Strep tococcus agalactiae, Strep. dysgalactiae, and Strep. uberis that targeted 16S–23S rDNA spacer regions. However, sensitivity of this multiplex PCR was less than for each individual PCR assay, and enrichment was needed to bring the threshold for detection by multiplex PCR for all bacteria assayed to 1 cfu/ml. A multiplex PCR assay for rapid and simultaneous detection of five foodborne pathogenic bacteria, including S. aureus, was developed by Kim and coworkers.75 Specific primers for multiplex PCR amplification of the Shiga-like toxin (verotoxin type II), femA (cytoplasmic protein), toxR (transmembrane DNA binding protein), iap (invasive associative protein), and invA (invasion protein A) genes were designed to allow simultaneous detection of Escherichia coli O157:H7, S. aureus, Vibrio parahaemolyticus, Listeria monocytogenes, and Salmonella, respectively. Finally, a new multiplex PCR-based procedure followed by capillary gel electrophoresis with laser-induced fluorescence detection (multiplex-PCR-CGE-LIF) was approached for the simultaneous detection of S. aureus, Listeria monocytogenes, and Salmonella spp.76 As compared to slab gel electrophoresis, the use of CGE-LIF improved the sensitivity of the multiplex PCR analysis by 10- to 1000-fold, allowing the detection of 2.6 × 103 cfu/ml of S. aureus, 570 cfu/ml of L. monocytogenes, and 790 cfu/ml of Salmonella in artificially inoculated food, without enrichment. Following 6 h of enrichment, as low as 260, 79, and 57 cfu/ml of S. aureus, L. monocytogenes, and Salmonella, respectively, were detected. The multiplex-PCR-CGE-LIF proved a powerful analytical tool to detect various food pathogens simultaneously in a fast, reproducible, and sensitive way. Even for rapid and reliable detection of S. aureus and its enterotoxins in food, a number of multiplex PCR (mPCR)based assays have been reported.33,77–80 In most of these studies, separate reactions are required to identify subsets of these genes. For example, two multiplex PCRs were developed for the detection of enterotoxigenic strains of S. aureus, one multiplex reaction for the simultaneous detection of enterotoxigenic strains type A (entA), type B (entB), and type E (entE) and another for the simultaneous detection of enterotoxigenic strains type C (entC) and type D (entD).79 These two multiplex PCRs were used to determine the presence of enterotoxigenic types for 51 S. aureus strains isolated from meat (sausage, ham, and chorizo) and dairy (powdered milk and cheese) products. Levels of correlation between the presence of genes that code for the production of SE (as determined by PCR) and the expression of these genes (as determined by the indirect enzyme-linked immunosorbent assay, ELISA) were 100% for SEA and SEE, 86% for SEC, 89% for SED, and 47% for SEB. Another multiplex PCR assay was described for the detection and differentiation of enterotoxigenic S. aureus in dairy products.18 In this case, also a solvent extraction procedure was successfully developed for extraction of S. aureus DNA from 10 ml of artificially contaminated skim milk or 20 g

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cheddar cheese. Primers targeting the enterotoxin C gene (entC) and thermostable nuclease gene (nuc) were used in the multiplex PCR. DNA was consistently quantified and amplified by uniplex PCR from 10 cfu/ml of S. aureus in skim milk or 10 cfu/20 g cheddar cheese. Even in this case, the sensitivity of the multiplex PCR was lower than for each individual PCR assay, resulting 100 cfu/ml of skim milk and 100 cfu/20 g cheddar cheese. Nevertheless, the developed methodology allows presumptive identification and differentiation of enterotoxigenic S. aureus in less than 6 h. A multiplex PCR for the simultaneous detection of S. aureus 23S rRNA, the coagulase and thermonuclease genes as well as the enterotoxin genes sea, sec, sed, seg, seh, sei, sej, sel was developed81 in order to obtain multiplex amplification products also starting from as little as 1 pg of DNA, corresponding approximately to 10 cfu/ml, showing the excellent specificity and high sensitivity of the assay. This multiplex PCR assay was used to correlate the distribution of genes encoding SEs with the presence of the corresponding SE production in S. aureus isolated from bovine, goat, sheep, and buffalo milk and dairy products.82 A total of 112 strains of S. aureus were tested for SE production by immuno-enzymatic (SEA–SEE) and reversed passive latex agglutination (SEA–SED) methods, while multiplex PCR was applied for SE genes (sea, sec, sed, seg, seh, sei, sej, and sel). Of the total strains studied, 67% were detected to have some SE genes (se), but only 52% produced a detectable amount of the classic antigenic SE types. The bovine isolates frequently had enterotoxin SEA, SED, and SEJ, while SEC and SEL predominated in the goat and sheep strains. The results demonstrated marked enterotoxigenic S. aureus strain variations, in accordance with strain origin and the two methods resulted in different information but concurred on the risk of foodstuff infection by S. aureus (Figure 18.1). 18.1.4.3 Real-Time PCR In recent years, an advanced form of PCR, real-time PCR, has been introduced into clinical microbiology. Compared with conventional PCR, real-time PCR is faster, more suited to high throughput of samples, and can quantitate the nucleic acid concentration. In real-time PCR, amplified products are detected by fluorescence at the moment that they are generated and directly related to the input target quantity, so that quantization is possible. The use of real-time PCR has driven significant changes in the microbial detection procedures. The predominantly phenotype-related methods of culture and antigen detection are being supplanted by the detection, characterization, and quantification of microbial nucleic acids. In comparison with conventional PCR, real-time PCR is more rapid, sensitive, reproducible and reduces the risk of carry-over contamination. The majority of real-time PCR applications in microbiology are for qualitative (resulting in a yes or no answer) detection of a virus, bacterium, fungus or parasite. For this purpose, fluorogenic PCR-based assays have shown promise in the detection of a variety of organisms including bacteria.83–85

Molecular Detection of Foodborne Pathogens

Real-time PCR utilizes the 5′–3′ nuclease activity of Taq DNA polymerase to digest an internal fluorogenic probe labelled with a fluorescent reporter dye and a fluorescent quencher dye.86 During amplification, the probe is hydrolyzed, relieving the quenching of the reporter dye, resulting in an increase in fluorescent intensity. This change in reporter dye fluorescence is quantitative for the PCR product, and under appropriate conditions, for the template. Such methods would be even more useful if they could be fine-tuned to simultaneously detect and quantify a mixture of pathogens in a sample. For example, a multiplex realtime PCR method to simultaneously detect common mastitis pathogens including S. aureus, Strep. agalactiae, and Strep. uberis directly from milk were developed by Gillespie and coworkers.52 This assay is the first to use a multiplex real-time PCR format for detection of pathogens directly from milk. Use of this method eliminates the need for post-PCR analysis by gel electrophoresis. This multiplex real-time PCR assay used information concerning primers for S. aureus and Strep. agalactiae that were shown to be sensitive and specific. Primers and probes for S. aureus were designed within the region used by Reischl et al.87 and Martineau et al.59 The cfb gene encoding the CAMP factor was used as the genetic target for this real-time PCR assay for detection of Strep. agalactiae. A strategy based on 5’ nuclease Triplex PCR was also developed for the rapid detection of nine enterotoxin genes (sea, seb, sec, sed, see, seg, seh, sei, sej) of S. aureus.10 This assay was first evaluated using a collection of S. aureus reference strains and then by testing previously characterized S. aureus strains isolated from food. While these assays correctly detected the SE genes in all the reference strains, in tests with field strains there was generally excellent agreement with the results obtained by conventional PCR, except for some strains harboring variant SE genes. The detection limits of these assays were also evaluated using fivefold dilution of recombinant plasmids for each se gene, ranging from 16 to 2000 copies of target se genes in the PCR tube. The development of this method is an improvement that should facilitate epidemiological investigations of SFP outbreaks. Two real-time fluorogenic PCR assays were described for the detection of entA, the gene that encodes SEA.88 The assays are useful in detecting and identifying strains of S. aureus that produce SEA and can serve a confirmatory role in determining the presence of SEA in food samples. The assays were tested in two real-time PCR formats, using either dye-labelled DNA probes corresponding to each primer set that are degraded by the 5’ exonuclease activity of Taq polymerase, or a PCR master mix that contains the DNA-binding dye SYBR Green. In both formats the assays have a limit of detection of between one and 13 copies of a S. aureus genome that contains a copy of entA. Neither assay crossreacted with genomic DNA isolated from other strains of S. aureus or other species. Two real-time quantitative PCR (RTQ-PCR) systems using nuc targeted primers, incorporating SYBR-Green I and TaqMan, respectively, have proven specific and suitable for the

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Staphylococcus

M

1

2

3

4

5

6

7

8

9

M

500 bp

Figure 18.1  Examples of multiplex PCR results for the detection of Staphylococcus aureus strains isolated from raw milk and raw cheese samples. Lanes 1–3, raw bovine milk samples; lane 4, curd, lanes 5–7 raw soft, semi-hard and hard cheese samples; lane 8, positive control; lane 10, negative control. All the samples were positive for 23S rRNA (499 bp), thermonuclease (nuc) (400 bp) and coagulase (coa) (204 bp) genes. The samples in lanes 4 and 5 are nonenterotoxigenic while the other samples contain se genes: lane 1, sample positive for sea gene (180 bp); lane 2, sample positive for sea (180 bp) sej (306 bp) and sed (343 bp) genes; lane 3, sample positive for sec (371 bp) gene; lane 6, sample positive for seg (432 bp) gene; lane 7, sample positive for sel (240 bp) and sec (371 bp) genes; lanes 8, reference strain ATCC 19095 (genes sec, seh, seg, sei, sel). M: 100 bp DNA ladder (Finnzymes).

routine detection and quantification of S. aureus in different food matrices.76 The two real-time PCR approaches improved the sensitivity of conventional PCR by decreasing the detection level to 10 (SYBR-Green I system) and 100 cells (TaqMan). In particular, the SYBR-Green I RTQ-PCR approach established allows the sensitive, automated and quantitative detection of S. aureus for routine analysis at a reasonable cost. A nuc targeted real-time PCR assay was developed to evaluate the risk associated with the enterotoxigenicity of S. aureus in Monte Veronese, a Protected Designation of Origin (PDO) cheese of the Lessinia area in Italy.89 By realtime quantification S. aureus numbers in cheese were found to exceed the limit tolerated by the Italian food legislation in 78% of the instances. Another real-time PCR detection system using a primer set from the sequence of the heat shock protein gene (hsp) and a gene coding for high-temperature-requirement A protein (htrA) was successfully developed to detect 16 S. aureus reference strains and 40 isolates from food-poisoning cases.90 When this primer set was used for the real-time PCR detection of S. aureus in milk and meat samples without the pre-enrichment step, samples with target cell numbers greater than 103 cfu/ml or cfu/g could be detected, indicating the potential quantitative ability of this real-time PCR assay. With a 10 h pre-enrichment step, however, a detection limit of 1 cfu/ml or cfu/g could be obtained. As the classical diagnostic bioassays of SEs as well as the routinely used immunological methods are hampered by several drawbacks regarding sensitivity, specificity, and practicability, sensitive reverse transcription-quantitative PCR

procedures can also be suitable for the routine detection and quantification of S. aureus and its enterotoxins in food. To analyze the expression of the enterotoxin genes of S. aureus, a reverse transcription real-time PCR was developed.91 Thanks to this assay, various enterotoxin genes were detected, including sea, seg, seh, sei, sen, seo, and sem. When the mRNA detection of the enterotoxin genes was analyzed using a reverse transcriptase PCR, various levels of expression were found depending on the species and enterotoxin gene. Therefore, it is reasonable to suggest that the poisoning risk of S. aureus can be effectively evaluated based on the gene expression at the mRNA level. In addition, a quantitative real-time immuno-PCR (qRT-iPCR) was developed for the detection of SEA and SEB and compared to a commercially available enzyme immunoassay.92 This qRT-iPCR approach was shown to overcome clearly the sensitivity limit of traditional immunological detection procedures for bacterial toxins, as demonstrated in this study for SEs. The development of a stable antibody-DNA conjugate providing a universal signal of amplification offers a versatile as well as a highly sensitive and specific tool for diagnostic and research purposes generally applicable for preformed antibody-antigen complexes. Finally, a quantitative PCR assay was also developed for the diagnosis of udder infections with S. aureus.93 For clinical milk samples, the analytical sensitivity of this assay was 50.7 times and 507 times greater than conventional bacteriology with 100 and 10 µl, respectively. Therefore, this assay, allowing the highly specific detection of S. aureus in bovine milk samples at very low concentrations, might become an important diagnostic tool.

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18.2  Methods 18.2.1 Reagents and Equipment Sample Treatment Reagents Isotonic diluent Alcohol Chlorexidine

Equipment Needed Pad Towels Cotton swabs Aluminum foil Sterile 10 ml tubes Sterile surgical blade Polyethylene stomacher bag

DNA Extraction Protocol Guanidine thiocyanate Sterile 1.5 ml microcentrifuge tubes EDTA Centrifuge for Eppendorf Tris-HCl Timer Triton X-100 Vortex DTT, Dithiothreitol Heat Eppendorf block 65°C Silica (Sigma Aldrich S5631) Pipette tips 1000 µl Absolute ethanol Isopropanol NaCl

Pipette tips 200 µl Disposable gloves

DNA Amplification Accuprime Taq DNA polymerase Sterile 0.2 ml microcentrifuge tubes (Invitrogen 12339-016) Double distilled water Thermal cycler Pipette tips 1000 µl Pipette tips 200 µl DNA molecular size marker 100 bp Pipette tips 20 µl DNA molecular size marker 1 kb Disposable gloves Ethidium bromide Centrifuge for Eppendorf Loading dye Gel electrophoresis apparatus Power supply Ultra violet source Agarose

18.2.2 Sample Collection and Preparation 18.2.2.1 Sample Collection General consideration. Depending upon the situation, the specimen may be collected from patients, controls, food handlers, animals, and food. The specimens should include the following: • Samples from patients and controls (e.g., serum, stool, vomitus, and urine) • Blood, spleen, intestinal content and liver tissue from fatal cases • Stool/rectal swabs, throat swabs and exudates/pus from lesions, if any, of food handlers • Sample of left over suspect food • Swabs of equipment/utensils with which food was processed • Samples from animals (e.g., milk) The general principles regarding collection of specimens are:

• Collect specimens aseptically with a sterile implement and collect in sterile containers. • Collect the specimens of the suspect food at the earliest. • If the food article is solid cut it with a sterile knife and collect 100–200 g of the sample from the centre. • In case of liquids, first thoroughly shake the specimen to mix and then with the help of a sterile tube collect the specimen and shift into a sterile container. • In case of raw meat or poultry, aseptically cut portions of meat/skin and put 100–200 g in a sterile plastic jar and refrigerate the specimen. • To collect the specimen from utensils and/or equipment in which food has been processed, moisten the swab with sterile 0.1% peptone water or buffered distilled water and swab the contact surfaces of the utensils. Then put the swab in an enrichment broth. • Collect aseptically a sample of the water, used for cooking purposes in a quantity ranging from 1 to 5 l as feasible. • All the specimens should be properly labeled and packed. The following details should accompany the specimens: • Place and address of location where the outbreak occurred • Symptoms: nausea/vomiting/diarrhea/fever/uncons­ ciousness, etc. • Date of onset of symptoms • Date specimens collected • Method of collection and transportation • Epidemiological background and the suspect organism • Condition of the food at the time of collection The specimen of food collected for analysis should be transported in sterile containers (plastic bag, tubes, etc.) to the laboratory by the most rapid mode available. Perishable food should be kept at 2–8oC. Hot food should be cooled rapidly by putting the containers under cold running water and then held at 0–4oC. Samples should be packed in such a way that there is no spillage during transportation. The receiving laboratory should be pre-informed about the method of transport and anticipated time of receipt in the laboratory. Milk sampling. Monthly bulk milk tank culturing has proven useful in monitoring udder health, particularly with regard to S. aureus.94 The sensitivity of a single bulk tank culture is fairly low, especially when the herd prevalence of S. aureus mastitis is low as well. In other words, often one bulk tank sample will be culture negative for S. aureus even though a herd may have cows infected with this organism. However, bulk tank cultures are highly specific (94%). So, it is rare that a bulk tank culture will be positive when in reality no cows in the herd have staphylococcal mastitis.

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Multiple sampling will improve the sensitivity of bulk tank culturing, particularly with intermittently shedding organisms like S. aureus. Serial testing can be performed by aseptically collecting an agitated bulk tank sample in a sterile container. This procedure can be repeated every other day when the bulk tank contains four milkings. The samples can be frozen immediately after they are obtained and delivered to the testing facility once each month. Milk cultures are the best method to determine if intramammary infection is due to S. aureus. A small (3–5 ml), sterile quarter sample is preferable to a voluminous contaminated composite. Teat ends should be thoroughly scrubbed with an alcohol pad or chlorexidine towels. The fore-milk should be discarded and a mid-stream milk sample obtained in a sterile container. Milk cultures should be immediately chilled to prevent overgrowth of environmental bacteria. If microbiologic procedures are to be delayed, the samples should be frozen. Meat sampling. The wet-dry sampling method95 involves the use of jumbo-head cotton swabs. Samples are collected within an area of 25 cm2 (2.5 × 10 cm). The sampling area is delimited by an autoclaved aluminium foil frame (2.5 ×   10 cm). For each sampling area, a swab is moistened in an isotonic diluent (peptone bacteriological, 1.0 g/l, and sodium chloride, 8.5 g/l) and then rubbed firmly across the sampling area with five strokes. This procedure is then repeated twice, with an angle of approximately 60° each time to ensure an even recovery of bacteria. The procedure is immediately repeated on the same area with a dry swab. Each pair of wet and dry swabs is combined into a single sample in a sterile universal test tube containing 10 ml of isotonic diluent and placed in the refrigerator until microbiological analyses are carried out. Excision-based sampling involves removing a sliver of tissue (2.5 × 10 cm, 3 mm thick) from a meat/skin section. An autoclaved aluminum foil frame (2.5 × 10 cm) is placed over the section, and an initial cut to a depth of approximately 4 mm is made with a sterile surgical blade. The same blade is then used to cut free the tissue sliver from the meat/skin section. Each sample is stored in a single, sealed, polyethylene stomacher bag and placed in the refrigerator for later microbiological analyses. 18.2.2.2 Sample Pretreatment A working method for rapid DNA extraction directly from bovine, ovine, and caprine raw milk and from fresh, soft, semi-hard and hard cheeses samples has been described.54,55 This method is based on the ability of silica-resin to bind DNA in the presence of a high concentration of guanidine thiocyanate chaotropic agent which guarantees an excellent disruption of bacterial cells from the main foodborne pathogens such as Staph. aureus. Milk samples. Dilute 500 µl of milk sample with 500 µl of sterilized saline solution (NaCl 0.9%) and centrifuge for 15 min at 600 × g at 4°C; discard the supernatant containing fat and liquid phase. Repeat the step once. Add to the pellet 50 µl of sterilized saline solution. Resuspend the pellet with vortex (20 sec). The solution will be turbid. Raw cheese samples. Dilute 100 mg of cheese sample with 500 µl of lysis buffer (3 M guanidine thiocyanate, 20

mM EDTA, 10 mM Tris–HCl, pH 6.8, 40 mg/l Triton X-100, 10 mg/l DTT) and vortexed for 30 sec to obtain an emulsified solution. 18.2.2.3 DNA Extraction (1) Add 300 µl of lysis buffer and 200 µl of binding solution (40 mg/ml silica from Sigma Aldrich, Milan, Italy, directly suspended in the lysis buffer) to the pellet previously resuspended in 50 µl of saline solution for milk sample or to the emulsified solution for cheese sample. Mix and incubate for 5 min at room temperature. Centrifuge for 30 sec at 450 × g (or 30 sec at 500 × g for cheese sample) and discard the supernatant. (2) Add 200 µl of lysis buffer and resuspend by vortexing (20 sec). Centrifuge for 30 sec at 450 × g (or 30 sec at 500 × g for cheese sample) and discard the supernatant. Repeat this step once. (3) Add 200 µl of washing solution (25% absolute ethanol, 25% isopropanol, 100 mM NaCl, 10 mM TrisHCl, pH 8) and resuspend by vortexing (20 sec). Centrifuge for 30 sec at 450 × g (or 30 sec at 500 × g for cheese sample) and discard the supernatant. Repeat this step once. (4) Add 200 µl of absolute ethanol solution and resuspend by vortexing (20 sec). Centrifuge for 30 sec at 450 × g (or 30 sec at 500 × g for cheese sample) and discard the supernatant. (5) Vacuum-dry the pellet in an Eppendorf heat block at 56°C for 10 min (or at room temperature for at least 15–20 min). (6) Add 100 µl of elution buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA), resuspend the pellet vortexing for 20 sec and incubate for 15 min at 65°C. (7) Centrifuge for 5 min at 450 × g (or 5 min at 500 × g for cheese sample) and transfer the supernatant, containing the DNA, into a clean tube. Note: to increase the DNA yield, a second elution step (with 5 min heating) may be performed. The solution contains pure DNA useful for molecular biology techniques. It can be used immediately or stored at –20°C for up to 6 months.

18.2.3  Detection Procedure In this section, a multiplex-PCR-based protocol is described81 with 11 primer sets to simultaneously identify the species together with two associated virulence marker genes (coagulase and thermostable nuclease genes) and eight of the known SE genes in S. aureus strains isolated from milk and dairy products. All primers used in the study, ranging from 20 to 24-mers, were designed using the Primer3 programme (http://wwwgenome.wi.mit.edu/cgibin/primer/primer3_www.cgi) except for the SEI forward primer taken from the literature.62

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Table 18.2 Primer Identities, Sequences, and Predicted Sizes of PCR Product for the Amplification of Staphylococcus aureus Target Genes Primer Identity 23S-F1200 23S-R1698 NUC-F166 NUC-R565 COA-F2591 COA-R2794 SEA-F1170 SEA-R1349 SEC-F97 SEC-R467 SED-F578 SED-R916 SEG-F322 SEG-R753 SEH-F260 SEH-R722 SEI-F71 SEI-R637 SEJ-F349 SEJ-R654 SEL-F158 SEL-R397

Primer Sequence (5′–3′) AGC TGT GGA TTG TCC TTT GG TCG CTC GCT CAC CTT AGA AT AGT TCA GCA AAT GCA TCA CA TAG CCA AGC CTT GAC GAA CT CCG CTT CAA CTT CAG CCT AC TTA GGT GCT ACA GGG GCA AT TAA GGA GGT GGT GCC TAT GG CAT CGA AAC CAG CCA AAG TT ACC AGA CCC TAT GCC AGA TG TCC CAT TAT CAA AGT GGT TTC C TCA ATT CAA AAG AAA TGG CTC A TTT TTC CGC GCT GTA TTT TT CCA CCT GTT GAA GGA AGA GG TGC AGA ACC ATC AAA CTC GT TCA CAT CAT ATG CGA AAG CAG TCG GAC AAT ATT TTT CTG ATC TTT CTC AAG GTG ATA TTG GTG TAG G CAG GCA GTC CAT CTC CTG TA GGT TTT CAA TGT TCT GGT GGT AAC CAA CGG TTC TTT TGA GG CAC CAG AAT CAC ACC GCT TA CTG TTT GAT GCT TGC CAT TG

Amplicon Size (bp)

Target Gene

499

23S rRNA 23S rRNA nuc nuc coa coa sea sea sec sec sed sed seg seg seh seh sei sei sej sej sel sel

400 204 180 371 339 432 463 529 306 240

Source: Cremonesi, P. et al. Development of a multiplex PCR assay for the identification of Staphylococcus aureus enterotoxigenic strains

isolated from milk and dairy products. Mol. Cell. Probes, 19, 299, 2005.

The primer sequences and the PCR product lengths are shown in Table 18.2. (1) In a 200 µl microtube, allocate 50 µl reaction mixture containing 2 µl DNA template, 2 U of AccuPrimeTM Taq DNA polymerase (Invitrogen,), 5 µl of 10 ×  AccuPrimeTM PCR buffer II containing 2 mM of each dNTPs (Invitrogen), 10 µM of the primer pair 23S-F1200 and 23S-R1698, 20 µM of the primer pairs COA-F2591 and COA-R2794, SEA-F1170 and SEA-R1349, SEC-F97 and SEC-R467, SED-F578 and SED-R916, SEJ-F349 and SEJ-R654, SEL-F158 and SEL-R397, 30 µM of the primer pairs NUCF166 and NUC-R565, SEG-F322 and SEG-R753, 40 µM of the primer pairs SEH-F260 and SEH-R722, SEI-F71 and SEI-R637; with double-distilled water to the final volume of 50 µl. (2) Perform PCR amplification in a thermal cycler GeneAmp PCR System 2700 (Applied Biosystems) with an initial denaturation at 94°C for 5 min, followed by 30 cycles of denaturation at 94°C for 1 min, primer annealing at 56°C for 1 min and extension at 68°C for 1 min, followed by a final extension at 72°C for 7 min. (3) Separate the amplified PCR products by 4% agarose gel electrophoresis (GellyPhor, Euroclone), and

include a DNA molecular size markers (100-bp and 1-kb DNA ladder; Finnzymes) in each agarose. (4) Stain the gel with ethidium bromide (0.05 µg/µl; Sigma Aldrich), and photograph the gel under ultra violet light using the BioProfile system (Mitsubishi).

18.3 Conclusions and Further Perspectives In conclusion, DNA-based methods to identify S. aureus in food have proved to be more sensitive and rapid than the conventional bacteriological methods. However, all these molecular methods should be standardized to become available for the routine analysis of food or clinical samples in laboratories. These systems are currently used only to confirm the diagnosis of specific bacterial strains. In addition, either a complex PCR with a mixture of large numbers of primers is needed, or a large series of individual PCRs must be run in parallel, or sequentially, to identify different bacterial strains (i.e., different enterotoxigenic S. aureus strains) or different pathogens contained in the same food or clinical sample. Recently, some of these problems were solved by using DNA microarrays. DNA microarrays are a promising diagnostic tool presenting many advantages compared to PCR or classical hybridization-based assays. They can detect tens of thousands

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of DNA sequences in a single hybridization step, allowing not only species determination but also a profiling of virulence factors genes. In the last decade, many DNA microarray have been developed for direct detection of the specific bacterial genes in clinical and food samples. Short oligonucleotide, PCR amplicons, whole genome or genome fragment microarrays have been successfully tested for detection of bacteria.93–112 Genetic variations in 16S rDNA could be utilized for the probe designing used in array identification system. With such a principle, Chiang and coworkers113 have developed a 16S rDNA-based oligonucleotide array system for the rapid diagnosis of the genus and species of bacteria in foods and clinical specimens. By using an array of specific oligonucleotides designed from the variable regions of 16S rDNA sequences, and the reverse hybridization of 16S rDNA amplified products to this array, the specific hybridization patterns for strains of Bacillus spp., E. coli, Salmonella spp., Staphylococcus spp. and Vibrio spp. were established. Another rapid and reliable one-tube microarray based assay was also developed for the simultaneous detection and identification (genetic typing) of almost all known enterotoxin genes of S. aureus.114 This method includes PCR amplification of part of the ent genes with universal primers, followed by analysis of amplicons by hybridization with ent-specific oligonucleotide probes immobilized on the microchip. More recently, Santini and coworkers115 have developed a prototype of medium density gene-segments DNA microarray for detection of the more prominent pathogens causing bloodstream infections, which includes the capture probes for S. aureus. All these DNA hybridization assays are suitable for S. aureus diagnosis; however faster, cheaper, miniaturized, multianalyte, easier-to-use, and more sensitive approaches are highly desired, especially in the case of decentralized analysis. In this context, electrochemical detection of DNA hybridization events offers innovative routes.116,117 Recently, the simultaneous detection of different food pathogenic bacteria by means of a disposable electrochemical low density genosensor array was described.118 The analytical method relied on the use of screen-printed arrays of gold electrodes, modified using thiol-tethered oligonucleotide probes. The samples identifying the bacteria of interest were obtained from the corresponding genomic DNAs through PCR amplification. These unmodified PCR products were captured at the electrode interface via sandwich hybridization with surface-tethered probes and biotinylated signalling probes. The resulting biotinylated hybrids were coupled with a streptavidin–alkaline phosphatase conjugate and then exposed to a α-naphthyl phosphate solution. Differential pulse voltammetry was finally used to detect the α-naphthol signal. The results of these studies demonstrate the usefulness of the microarray assay for the analysis of food pathogens in general and for the analysis of multitoxigenic strains, such as S. aureus strains. Microarrays are not in common use in average laboratories today. However, like any new technology, as more applications are developed for the microarray

technology, it will become more practical and may well become widely used.

References









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Molecular Detection of Foodborne Pathogens 38. Ramesh, A. et al. Application of a convenient DNA extraction method and multiplex PCR for the direct detection of Staphylococcus aureus and Yersinia enterocolitica in milk samples. Mol. Cell. Probes, 16, 307, 2002. 39. Rossen, L. Inhibition of PCR by components of food samples, microbial diagnostic assays and DNA-extraction solutions. Int. J. Food Microbiol., 17, 37, 1992. 40. Wilson, I.G. Minireview: inhibition and facilitation of nucleic acid amplification. Appl. Environ. Microbiol., 63, 3741, 1997. 41. Straub, J.A., Hertel, C. and Hammes, W.P. A 23S rDNA-targeted polymerase chain reaction-based system for detection of Staphylococcus aureus in meat starter cultures and dairy products. J. Food Prot., 62, 1150, 1999. 42. Hein, I. et al. Real-time PCR for the detection of Salmonella spp. in food: an alternative approach to a conventional PCR system suggested by the FOOD-PCR project. J. Microbiol. Methods, 66, 538, 2006. 43. Yang, S. et al. Quantitative multiprobe PCR assay for simultaneous detection and identification to species level of bacterial pathogens. J. Clin. Microbiol., 40, 3449, 2002. 44. Jung, Y.S., Frank, J.F. and Brackett, R.E. Evaluation of antibodies for immunomagnetic separation combined with flow cytometry detection of Listeria monocytogenes. J. Food Prot., 66, 1283, 2003. 45. Lantz, P.G. et al. Enhanced sensitivity in PCR detection of Listeria monocytogenes in soft cheese through use of an aqueous two-phase system as a sample preparation method. Appl. Environ. Microbiol., 60, 3416, 1994. 46. Lindqvist, R., Norling, B. and Lambertz, S.T. A rapid sample preparation method for PCR detection of food pathogens based on buoyant density centrifugation. Lett. Appl. Microbiol., 24, 306, 1997. 47. Wolffs, P. et al. Rapid quantification of Yersinia enterocolitica in pork samples by a novel sample preparation method, flotation, prior to real-time PCR. J. Clin. Microbiol., 42, 1042, 2004. 48. Allmann, M. et al. Polymerase chain reaction (PCR) for detection of pathogenic microorganisms in bacteriological monitoring of dairy products. Res. Microbiol., 146, 85, 1995. 49. Wegmuller, B., Luthy, J. and Candrian, U. Direct polymerase chain reaction detection of Campylobacter jejuni and Campylobacter coli in raw milk and dairy products. Appl. Environ. Microbiol., 59, 2161, 1993. 50. Martinez, G., Harel, J. and Gottschalk, M. Specific detection by PCR of Streptococcus agalactiae in milk. Can. J. Vet. Res., 65, 68, 2001. 51. Hein, I. et al. Detection and quantification of the iap gene of Listeria monocytogenes and Listeria innocua by a new realtime quantitative PCR assay. Res. Microbiol., 152, 37, 2001. 52. Gillespie, B.E. and Oliver S.P. Simultaneous detection of mastitis pathogens, Staphylococcus aureus, Streptococcus uberis, and Streptococcus agalactiae by multiplex real-time polymerase chain reaction. J. Dairy Sci., 88, 3510, 2005. 53. Fukushima, H. et al. Rapid separation and concentration of food-borne pathogens in food samples prior to quantification by viable-cell counting and real-time PCR. Appl. Environ. Microbiol., 73, 92, 2006. 54. Cremonesi, P. et al. Technical note: improved method for rapid DNA extraction of mastitis pathogens directly from milk. J. Dairy Sci., 89, 163, 2006. 55. Cremonesi, P. et al. Detection of enterotoxigenic Staphylococcus aureus isolates in raw milk cheese. Lett. Appl. Microbiol., 45, 586, 2007.

Staphylococcus 56. Kubota, M. et al. Rapid and effective method for separation of Staphylococcus aureus from somatic cells in mastitis milk. J. Dairy Sci., 90, 4100, 2007. 57. Johnson, W.M. et al. Detection of genes for enterotoxins, exfoliative toxins, and toxic shock syndrome toxin 1 in Staphylococcus aureus by the polymerase chain reaction. J. Clin. Microbiol., 29, 426, 1991. 58. Brakstad, O.G., Aasbakk, K. and Maeland, J.A., 1992. Detection of Staphylococcus aureus by polymerase chain reaction amplification of the nuc gene. J. Clin. Microbiol., 30, 1654, 1992. 59. Martineau, F. et al. Species-specific and ubiquitous-DNAbased assays for rapid identification of Staphylococcus aureus. J. Clin. Microbiol., 36, 618, 1998. 60. Atanassova, V. et al. Prevalence of Staphylococcus aureus and staphylococcal enterotoxins in raw pork and uncooked smoked ham-a comparison of classical culturing detection and RFLP-PCR. Int. J. Food Microbiol., 68, 105, 2001. 61. Amann, R.L., Ludwig, W. and Schleifer, K.H. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol. Rev., 59, 143, 1995. 62. Akineden, O. et al. Toxin genes and other characteristics of Staphylococcus aureus isolates from milk of cows with mastitis. Clin. Diagn. Lab. Immunol., 8, 959, 2001. 63. Chotar, M., Vidova, B. and Godany, A. Development of specific and rapid detection of bacterial pathogens in dairy products by PCR. Folia Microbiol. (Praha), 51, 639, 2006. 64. Riyaz-Ul-Hassan, S., Verma, V. and Qazi, G.N. Evaluation of three different molecular markers for the detection of Staphylococcus aureus by polymerase chain reaction. Food Microbiol., 25, 452, 2008. 65. Liu, Z.M., Shi, X.M., and Pan, F. Species-specific diagnostic marker for rapid identification of Staphylococcus aureus. Diagn. Microbiol. Infect. Dis., 59, 379, 2007. 66. Su, C. et al. Phylogenetic relationships of Staphylococcus aureus from bovine mastitis based on coagulase gene polymorphism. Vet. Microbiol., 71, 53, 2000. 67. Schlegelova, J. et al. Staphylococcus aureus isolates from dairy cows and humans on a farm differ in coagulase genotype. Vet. Microbiol., 92, 327, 2003. 68. Carter, P.E., Begbie, K. and Thomson-Carter, F.M. Coagulase gene variants associated with distinct populations of Staphylococcus aureus. Epidemiol. Infect., 130, 207, 2003. 69. Barski, P. et al. Rapid assay for detection of methicillin-resistant Staphylococcus aureus using multiplex PCR. Mol. Cell. Probes, 10, 471, 1996. 70. Kalorey, D.R. et al. PCR-based detection of genes encoding virulence determinants in Staphylococcus aureus from bovine subclinical mastitis cases. J. Vet. Sci., 8, 151, 2007. 71. Chiang, Y.C. et al. PCR detection of Staphylococcal enterotoxins (SEs) N, O, P, Q, R, U, and survey of SE types in Staphylococcus aureus isolates from food-poisoning cases in Taiwan. Int. J. Food. Microbiol., 121, 6, 2008. 72. Blaiotta, G. et al. PCR detection of staphylococcal enterotoxin genes in Staphylococcus spp. Strains isolated from meat and dairy products. Evidence for new variants of seG and seI in S. aureus AB-8802. J. Appl. Microbiol., 97, 719, 2004. 73. Henegariu, O. et al. Multiplex PCR-critical parameters and step-by-step protocol. Biotech., 23, 504, 1997. 74. Atlas, R.M. and Bej, A. K. 1994. Polymerase chain reaction. In: Gerhardt, P., Murray, R.G.E., Wood, W.A. and Krieg, N.R. (eds). Methods for General and Molecular Bacteriology. American Society for Microbiology, Washington, DC, 1994, 418.

257 75. Kim, L. et al. A novel multiplex PCR assay for rapid and simultaneous detection of five pathogenic bacteria: Escherichia coli O157:H7, Salmonella, Staphylococcus aureus, Listeria monocytogenes, and Vibrio parahaemolyticus. J. Food Prot., 70, 656, 2007. 76. Alarcon, B. et al. Simultaneous and sensitive detection of three foodborne pathogens by multiplex PCR, capillary gel electrophoresis, and laser-induced fluorescence. J. Agric. Food Chem., 52, 7180, 2004. 77. Monday, S.R. and Bohach, G.A. Use of multiplex PCR to detect classical and newly described pyrogenic toxin genes in staphylococcal isolates. J. Clin. Microbiol., 37, 3411, 1999. 78. Sharma, N.K., Rees, C.D. and Dodd, C.E.R. Development of a single-reaction multiplex PCR toxin typing assay for Staphylococcus aureus strains. Appl. Environ. Microbiol., 66, 1347, 2000. 79. Najera-Sanchez, G. et al. Development of two multiplex polymerase chain reactions for the detection of enterotoxigenic strains of Staphylococcus aureus isolated from foods. J. Food Prot., 66, 1055, 2003. 80. Rosec, J.P. and Gigaud, O. Staphylococcal enterotoxin genes of classical and new types detected by PCR in France. Int. J. Food Microbiol., 77, 61, 70, 2002. 81. Cremonesi, P. et al. Development of a multiplex PCR assay for the identification of Staphylococcus aureus enterotoxigenic strains isolated from milk and dairy products. Mol. Cell. Probes, 19, 299, 2005. 82. Morandi, S. et al. Detection of classical enterotoxins and identification of enterotoxin genes in Staphylococcus aureus from milk and dairy products. Vet. Microbiol., 124, 66, 2007. 83. Jourdan, A.D., Johnson, S.C. and Wesley, I.V. Development of a fluorogenic 5’ nuclease PCR assay for detection of the ail gene of pathogenic Yersinia enterocolitica. Appl Environ. Microbiol., 66, 3750, 2000. 84. Sharma, V.K. and Carlson, S.A. Simultaneous detection of Salmonella strains and Escherichia coli O157:H7 with fluorogenic PCR and single-enrichment-broth culture. Appl. Environ. Microbiol., 66, 5472, 2000. 85. Yoshida, A. et al. Development of a 5′ fluorogenic nuclease-based real-time PCR assay for quantitative detection of Actinobacillus actinomycetemcomitans and Porphyromonas gingivalis. J. Clin. Microbiol., 41, 863, 2003. 86. Cai, H.Y. et al. Molecular genetic methods in the veterinary clinical bacteriology laboratory: Current usage and future applications. Anim. Health Res. Rev., 4, 73, 2003. 87. Reischl, U. et al. Rapid identification of methicillin-resistant Staphylococcus aureus and simultaneous species confirmation using real-time fluorescence PCR. J. Clin. Microbiol., 38, 2429, 2000. 88. Horsmon, J.R. et al. Real-time fluorogenic PCR assays for the detection of entA, the gene encoding staphylococcal enterotoxin A. Biotechnol. Lett., 28, 823, 2006. 89. Poli, A. et al. Detection of Staphylococcus aureus and enterotoxin genotype diversity in Monte Veronese, a Protected Designation of Origin Italian cheese. Lett. Appl.Microbiol., 45, 529, 2007. 90. Chiang, Y.C. et al. Real-time PCR detection of Staphylococcus aureus in milk and meat using new primers designed from the heat shock protein gene htrA sequence. J. Food Prot., 70, 2855, 2007. 91. Lee, Y.D. et al. Expression of enterotoxin genes in Staphylococcus aureus isolates based on mRNA analysis. J. Microbiol. Biotechnol., 17, 461, 2007.

258 92. Fischer, A. et al. A quantitative real-time immuno-PCR approach for detection of staphylococcal enterotoxins. J. Mol. Med., 85, 461, 2007. 93. Graber, H.U. et al. Development of a highly sensitive and specific assay to detect Staphylococcus aureus in bovine mastitic milk. J. Dairy Sci., 90, 4661, 2007. 94. Harmon, R.J. et al. Microbiological Procedures for the Diagnosis of Bovine Udder Infection, 3rd Edn. National Mastitis Council, Inc., Arlington, VA, 1990. 95. Cenci-Goga, B.T. et al. An in vitro system for the comparison of excision and wet-dry swabbing for microbiological sampling of beef carcasses. J. Food Prot., 70, 930, 2007. 96. Laberge, I. et al. Detection of Cryptosporidium parvum in raw milk by PCR and oligonucleotide probe hybridization. Appl. Environ. Microbiol., 62, 3259, 1996. 97. Chizhikov, V. et al. Microarray analysis of microbial virulence factors. Appl. Environ. Microbiol., 67, 3258, 2001. 98. Hong, B.X. et al. Application of oligonucleotide array technology for the rapid detection of pathogenic bacteria of foodborne infections. J. Microbiol. Methods, 58, 403, 2004. 99. Zhang, H. et al. An electronic DNA microarray technique for detection and differentiation of viable Campylobacter species. Analyst, 131, 907, 2006. 100. Mozola, M.A. Genetics-based methods for detection of Salmonella spp. in foods. J. AOAC Int., 89, 517, 2006. 101. Myers, K.M., Gaba, J. and Al-Khaldi, S.F. Molecular identification of Yersinia enterocolitica isolated from pasteurized whole milk using DNA microarray chip hybridization. Mol. Cell. Probes, 20, 71, 2006. 102. Wang, X.W. et al. Development and application of an oligonucleotide microarray for the detection of food-borne bacterial pathogens. Appl. Microbiol. Biotechnol., 76, 225, 2007. 103. DeRisi, J.L., Iyer, V.R. and Brown, P.O. Exploring the metabolic and genetic control of gene expression on a genomic scale. Science, 278, 680, 1997. 104. Lashkari, D.A. et al. Yeast microarrays for genome wide parallel genetic and gene expression analysis. Proc. Natl. Acad. Sci. USA, 94, 13057, 1997. 105. Duggan, D.J. et al. Expression profiling using cDNA microarrays. Nat. Genet., 21, 10, 1999. 106. Cho, J.C. and Tiedje, J.M. Bacterial species determination from DNA–DNA hybridization by using genome fragments and DNA microarrays. Appl. Environ. Microbiol., 67, 3677, 2001. 107. Volokhov, D. et al. Identification of Listeria species by microarray-based assay. J. Clin. Microbiol., 40, 4720, 2002. 108. Volokhov, D. et al. Microarraybased identification of thermophilic Campylobacter jejuni, C. coli, C. lari, and C. upsaliensis. J. Clin. Microbiol., 41, 4071, 2003. 109. Volokhov, D. et al. Microarray analysis of erythromycin resistance determinants. J. Appl. Microbiol., 95, 787, 2003. 110. Bekal, S. et al. Rapid identification of Escherichia coli pathotypes by virulence gene detection with DNA microarrays. J. Clin. Microbiol., 41, 2113, 2003. 111. Wu, L. et al. Development and evaluation of microarray-based wholegenome hybridization for detection of microorganisms within the context of environmental applications. Environ. Sci. Technol., 38, 6775, 2004. 112. Vianna, M.E. Microarrays complement culture methods for identification of bacteria in endodontic infections. Oral Microbiol. Immunol., 20, 253, 2005. 113. Chiang, Y.C. et al. Identification of Bacillus spp., Escherichia coli, Salmonella spp., Staphylococcus spp. and Vibrio spp. with 16S ribosomal DNA-based oligonucleotide array hybridization. Int. J. Food Microbiol., 107, 131, 2006.

Molecular Detection of Foodborne Pathogens 114. Seergev, N. et al. Simultaneous analysis of multiple staphylococcal enterotoxin genes by an oligonucleotide microarray assay. J. Clin. Microbiol., 42, 2134, 2004. 115. Santini, M.P. et al. Rapid identification, virulence analysis and resistance profiling of Staphylococcus aureus by gene segment-based DNA microarrays: application to blood culture post-processing. J. Microbiol. Methods, 68, 468, 2007. 116. Lucarelli, F. et al. Carbon and gold electrodes as electrochemical transducers for DNA hybridisation sensors. Biosens. Bioelectron., 19, 515, 2004. 117. Kerman, K., Kobayashi, M. and Tamiya, E. Electrochemical molecular analysis without nucleic acid amplification. Meas. Sci. Technol., 15, R1, 2004. 118. Farabullini, F. et al. Disposable electrochemical genosensor for the simultaneous analysis of different bacterial food contaminants. Biosen. Bioelec., 22, 1154, 2007. 119. Betley, M.J. and Mekalanos, J.J. Nucleotide sequence of the type A staphylococcal enterotoxin gene. J. Bacteriol., 170, 34, 1988. 120. Jones, C.L. and Khan, S.A. Nucleotide sequence of the enterotoxin B gene from Staphylococcus aureus. J. Bacteriol, 166, 29, 1988. 121. Bohach, G.A. and Schlievert, P.M. Nucleotide sequence of the staphylococcal enterotoxin C1 gene and relatedness to other pyrogenic toxins. Mol. Gen. Genet., 209, 15, 1987. 122. Bohach, G.A. and Selievert, P.M. Conservation of the biologically active portions of staphylococcal enterotoxins C1 and C2. Infect. Immun., 57, 2249, 1989. 123. Hovde, C.J., Hackett, S.P. and Bohach, G.A. Nucleotide sequence of the staphylococcal enterotoxin C3 gene: sequence comparison of all three type C staphylococcal enterotoxins. Mol. Gen. Genet., 220, 329, 1990. 124. Marr, J.C. Characterization of novel type C staphylococcal enterotoxins: biological and evolutionary implications. Infect. Immun., 61, 4254, 1993. 125. Bayles, K.W. and Jandolo, J.J. Genetic and molecular analyses of the gene encoding staphylococcal enterotoxin D. J. Bacteriol., 171, 4799, 1989. 126. Couch, J.L., Soltis, M.T. and Betley, M.J. Cloning and nucleotide sequence of the type E staphylococcal enterotoxin gene. J. Bacteriol., 170, 2954, 1988. 127. Munson, S.H. et al. Identification and characterization of staphylococcal enterotoxin types G and I from Staphylococcus aureus. Infect. Immun., 66, 3337, 1998. 128. Su, Y.C. and Lee Wong, A.C. Identification and purification of a new staphylococcal enterotoxin, H. Appl. Environ. Microbiol., 61, 1438, 1998. 129. Zhang, S., Iandolo, J.J. and Stewart, G.C. The enterotoxin D plasmid of Staphylococcus aureus encodes a second enterotoxin determinant (sej). FEMS Microbiol. Lett., 168, 1998. 130. Orwin, P.M. et al. Biochemical and biological properties of Staphylococcal enterotoxin K. Infect. Immun., 69, 360, 2001. 131. Fitzgerald, J.R. et al. Characterization of a putative pathogenicity island from bovine Staphylococcus aureus encoding multiple superantigens. J. Bacteriol., 183, 63, 2001. 132. Omoe, K. et al. Characterization of novel staphylococcal enterotoxin-like toxin type P. Infect. Immun., 73, 5540, 2005. 133. Orwin, P.M. et al. Characterization of a novel staphylococcal enterotoxin-like superantigen, a member of the group V subfamily of pyrogenic toxins. Biochemistry, 41, 14033, 2002. 134. Omoe, K. et al. Identification and characterization of a new staphylococcal enterotoxin-related putative toxin encoded by two kinds of plasmids. Infect. Immun., 71, 6088, 2003. 135. Letertre, C. et al. Identification of a new putative enterotoxin SEU encoded by the egc cluster of Staphylococcus aureus. J. Appl. Microbiol., 95, 38, 2003.

19 Streptococcus Mark van der Linden

Institute of Medical Microbiology and National Reference Center for Streptococci

Romney S. Haylett

Institute of Immunology

Ralf René Reinert

Wyeth Vaccines Research

Lothar Rink

Institute of Immunology

Contents 19.1 Introduction.................................................................................................................................................................... 259 19.1.1 Classification..................................................................................................................................................... 259 19.1.2 Association of Streptococci with Foodborne Disease...................................................................................... 261 19.1.3 Pathogenesis...................................................................................................................................................... 261 19.1.4 Diagnosis........................................................................................................................................................... 261 19.1.4.1 Conventional Techniques................................................................................................................. 261 19.1.4.2 Molecular Techniques...................................................................................................................... 265 19.2 Methods.......................................................................................................................................................................... 265 19.2.1 Sample Collection and Preparation.................................................................................................................. 265 19.2.2 Detection Procedures........................................................................................................................................ 265 19.2.2.1 PCR for emm-Typing Group A Streptococci (GAS)........................................................................ 265 19.2.2.2 Multiplex PCR for the Detection of Group A Streptococcal Superantigen..................................... 265 19.3 Conclusion and Future Perspectives............................................................................................................................... 267 References.................................................................................................................................................................................. 267

19.1  Introduction

19.1.1 Classification

Diseases caused by bacteria of the genus Streptococcus are among the infections most often encountered by general practitioners and clinical doctors. Many of the diseases like streptococcal tonsillopharyngitis can be treated with antibiotics, whereas others such as meningitis and pneumonia caused by Streptococcus pneumoniae can be prevented by vaccination. Some other less common streptococcal infections are more difficult to treat including puerperal sepsis, which is caused by Streptococcus agalactiae, and streptococcal toxic shock-like syndrome (STSS). STSS is an illness associated with streptococcal superantigens (SAg), which can be typed by PCR (see Section 19.2.2). Rising antibiotic resistance rates, especially in Streptococcus pneumoniae, poses another large problem in the treatment of pneumococcal disease.1

The genus Streptococcus consists of 31 species.2 Several of these species are well known human (e.g., S. pneumoniae, S. pyogenes, S. agalactiae) or animal (e.g., S. bovis, S. canis, S. equinus, S. iniae) pathogens. Many streptococci belong to the commensal flora and only reveal their pathogenicity when the general condition of the host is weakened. Streptococci (Greek: pearls on a chain) are Gram-positive spherical bacteria that grow in chains, sometimes in pairs (S. pneumoniae, also called diplococcus). They are mostly grown on media containing sterile sheep blood under enhanced CO2 conditions. Most streptococci are facultative anaerobes. Streptococci can be classified according to their type of hemolyis. Hemolysis is the enzymatic breakdown of red blood cells that can be observed in blood agar plates. Streptococci show three types of hemolysis: α, β, and γ. α-hemolytic streptococci,

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like S. pneumoniae, show a greenish halo around the colonies, resulting from the reduction of hemoglobin to methemoglobin in the red blood cells. In β-hemolysis, the red blood cells are completely lysed, resulting in a clear halo around the colonies. β-hemolysing streptococci include S. pyogenes and S. agalactiae. γ-hemolysis is the term used to indicate no hemolysis at all where no discoloration of the blood can be observed. A serological classification system for streptococci was designed by Rebecca Lancefield and is based on specific carbohydrate antigens contained in the cell wall.3 The system divides the streptococci into groups A through T. Lancefield antigens can nowadays be determined using monoclonal antibodies in a simple agglutination test (see Section 19.2.2). Group A streptococci (GAS) are β-hemolytic streptococci, which contains, with very rare exceptions,4 only one species: S. pyogenes. GAS is a strictly human pathogen and infections are the most frequent especially among children. Infections range from respiratory tract and skin infections (tonsillopharyngitis, scarlet fever) to invasive life-threatening diseases like necrotizing fasciitis and STSS. Additionally, a number of sequelae such as acute rheumatic fever (ARF) and acute glomerulonephritis (AGN) are associated with GAS infections. For further classification, GAS can be subdivided into M-types, based on the presence of M-proteins on the cell surface.5 M-typing was previously performed with specific antibodies. Nowadays it is possible to determine the variable part of the emm-gene coding for the M-protein, resulting in so-called emm-types (see Section 19.2.2). The main member of the Group B streptococci (GBS) is S. agalactiae, which is commonly found as a member of the normal flora of the human vagina and urethra. GBS was first found among cows with chronic mastitis. S. agalactiae is one of the most important causes of neonatal infections.6 The newborns become infected during the first week of life when the pathogen is transferred from the mother to the child. Disease may develop within a few hours after birth (early onset), or between 4 days to several months later (late-onset). Typically, early onset disease includes severe sepsis and pneumonia, whereas late-onset disease mainly manifests itself as meningitis. S. agalactiae also belongs to the β-hemolytic streptococci. Serologically S. agalactiae can be subdivided into eight serotypes, based on the capsular polysaccharide. Serotypes can be determined using specific antisera in an agglutination test. Streptococci of the serological groups C and G are both human and animal pathogens. Human pathogens from these groups are comprised of S. dysgalactiae ssp. dysgalactiae and S. dysgalactiae ssp. equisimilis. Sometimes, these strains can be transferred from animals to humans (zoonosis).7 Groups D, E, and F streptococci may occasionally cause infections in humans, but are not as frequently associated with human diseases like groups A, B, C, and G streptococci. Until 20 years ago, Group D streptococci contained the fecal streptococci, which are now classified as enterococci. The only streptococcal species remaining in this group are S. bovis and S. equinus.8 Group E streptococci are mostly associated with infections in animals (cows, pigs).9 Group F streptococci make up part of the commensal flora of the human throat, and may sometimes cause infections (e.g., tooth infections).

Molecular Detection of Foodborne Pathogens

Streptococci of other Lancefield groups are rarely associated with human infections.10 S. pneumoniae is not classifiable with the Lancefield system. It is an α-hemolytic Streptococcus requiring blood and elevated CO2 levels for growth. It usually grows in pairs, and was therefore previously called Diplococcus pneumoniae. It is also referred to as pneumococcus. Pneumococci belong to the normal oropharyngal flora of many humans. About 50% of (young) children are colonized. Pneumococcal infections are predominantly found in young children (<5 years of age) followed by elderly (>60 years of age) and immunocompromised patients. Infections range from otitis media (mainly in children), to severe invasive infections like pneumonia, bacteraemia and meningitis.11 Pneumococci carry a polysaccharide capsule which helps evade the human immune defense. Currently, 91 different capsular types, called serotypes, have been identified. Determination of the serotype is done with the help of specific antibodies in a test called Neufelds Quellung reaction. The antibodies bind to the capsular polysaccharide, producing a swelling which can be detected under the microscope. The genes needed for the synthesis of the capsular polysaccharide are organized in a “cassette.” Recently the cassettes for all 91 serotypes were sequenced.12 This has opened opportunities for PCR based serotyping methods.13 An analysis of isolates from invasive pneumococcal disease (IPD) in the USA showed that seven serotypes were responsible for over 80% of IPD cases in young children.14 Analyses of IPD in other countries showed the serotype distribution to be different from region to region. Pneumococci display a typically clonal behaviour, with certain clones rising and falling over time. Usually, this is associated with drug resistance, i.e., clones that have acquired resistance to certain antibiotics and can now successfully spread among the population. Pneumococci are naturally competent organisms and can therefore take up DNA from their environment. This results in high levels of recombination even crossing species borders.15 It was shown that pneumococci acquired β-lactam resistance through recombination of DNA encoding penicillin-binding proteins (PBPs), from strains like S. mitis and S. oralis, resulting in so-called mosaic genes.16 The resistance situation of pneumococci is indeed worrisome. High levels of both β-lactam and macrolide resistance have been reported from many parts of the world, although there are large differences between countries. Resistance levels in Spain and France have reached levels of up to 50%, whereas levels in Germany (6%) and the Netherlands (<2%) are much lower.17,18 As mentioned above, the spread of resistance is often caused by the successful proliferation of certain clones. Currently, two vaccines exist against pneumococci. One is a 23-valent polysaccharide vaccine, which provides protection against the 23 serotypes most commonly found in infections. However, this vaccine does not elicit an immune response in children under 2 years of age. Recently, a seven-valent conjugate vaccine has been introduced, which does generate an immune response in children. A vaccination program using this vaccine has been successfully implemented in the United States. Recently Germany and many other countries worldwide have also started vaccination programs.

261

Streptococcus

19.1.2 Association of Streptococci with Foodborne Disease Before the introduction of conservation methods like pasteurization and refrigeration, foodborne infections caused by streptococci were very common. In more recent times they have become rarer due to better hygiene and improved conservation methods. Most current reports of outbreaks of foodborne streptoccocal infections are associated with group events (parties, conventions, etc.) or with situations where people live in close contact with each other (military bases, prisons, etc.).19 Outbreaks almost exclusively involve S. pyogenes (GAS), although some rare cases involving group G streptococci have been reported.20 An extensive overview of reports of foodborne infections in the past 60 years is given in Table 19.1. Foodborne outbreaks are generally associated with poor hygiene during food preparation. Use of unclean kitchen tools (knives etc.), not wearing gloves, dirty aprons, but also poor refrigeration of previously prepared food are often at the basis of outbreaks. The use of untrained personnel or volunteers not familiar with standard hygienic procedures also plays a role. Secondary spread is an important factor in situations where people live in close contact with each other such as on military bases and in prisons. Very often it is not possible to find the source of an outbreak due to the fact that the contaminated material, or the persons involved are not present anymore. However, since GAS are almost exclusive human pathogens, the sources are mostly human. Since foodborne infections caused by streptococci have become so rare, they often go unnoticed. Therefore, their suspected number is expected to be larger than reported. Furthermore, it must be assumed that foodborne infections with S. pyogenes are still prevalent in developing countries.19

19.1.3  Pathogenesis GAS disease belongs to the most prevalent infectious ­diseases worldwide, especially among children, and varies from mild skin infections to severe invasive diseases. The most common disease is tonsillopharyngitis.57 Another mild form of GAS disease is impetigo, a localized skin infection. Penetration of GAS into the deeper skin layers leads to erysipelas and cellulitis. Invasion of GAS into the blood leads to necrotizing fasciitis, a life-threatening disease. Scarlet fever is a special form of streptococcal pharyngitis caused by streptococcal pyrogenic exotoxins. Symptoms are fever, sore throat, strawberry tongue, and a fine rash over the entire body. The exotoxins belong to the so-called SAg. Release of exotoxins from invasive GAS may lead to STSS, another life-threatening disease. S. pyogenes can cause post-­infectious non-pyogenic sequelae. The most important ones are ARF and AGN. These conditions appear several weeks after the initial infection. GAS have several virulence factors. The capsule, composed of hyaluronic acid, protects cells from phagocytosis. M-protein present on the cell surface prevents opsonization. Attachment to host cells is facilitated by M-protein, lipoteichoic acid, and protein F. A number of virulence factors are released into the host. Streptolysins O and F are toxins responsible for β-hemolysis. Streptokinase is an enzyme that activates plasminogen into

plasmin, which digests fibrin and other proteins. Hyaluronidase facilitates the spread of GAS through connective tissue. A total of 11 SAg are secreted by GAS. The streptococcal pyogenic exotxins SPEA and SPEC are responsible for the rash of scarlet fever and many of the symptoms of streptococcal toxic shock syndrome. Other SAg are SPEG, SPEH, SPEI, SPEJ, SPEK, SPEL, SPEM, SSA, and SMEZ. Streptodornase is a mixture of up to four DNases secreted by GAS, protecting the cells from being trapped in neutrophil extracellular traps. One of the DNases was erroneously defined as SPEF, but is definitely not a superantigen and possesses no mitogenic activity. SPEB was also initially described as a superantigen, but was shown to be a cysteine proteinase which possesses mitogenic activity. C5a peptidase cleaves the neutrophil C5a and minimizes the influx of ­neutrophils in early infection. The serine protease ScpC prevents the migration of neutrophils to the spreading infection by degrading the chemokine IL-8. So far no penicillin resistant GAS have been reported, and thus penicillin remains the antibiotic of choice for the treatment of GAS disease. S. agalactiae is a species of the normal flora of the gut and female urogenital tract. Perinatal infections cause septicemia, pneumonia or meningitis, which are associated with a high mortality. S. agalactiae is one of the most important causes of neonatal infections like sepsis, meningitis, and pneumonia.6 The pathogen is transferred from mother to child during the first weeks of life. Disease may develop within a few hours after birth (early onset), or between 4 days to several months later (late-onset). Typically, early onset disease includes severe sepsis and pneumonia, whereas late-onset disease mainly manifests itself as meningitis. The capsular polysaccharide of GBS is the main virulence factor. S. pneumoniae colonizes the human nasopharynx. Asymptomatic carriage of pneumococci is found among 30–50% of all children and 5–10% of adults. Following colonization in the nasopharynx the bacteria can get carried to the Eustachian tube or nasal sinuses and cause otitis media and sinusitis. When pneumococci penetrate the mucous membrane and enter the lower respiratory tract they can cause pneumonia. Invasive S. pneumoniae can result in pneumococcal meningitis. Pneumococci evade phagocytosis through their polysaccharide capsule.

19.1.4  Diagnosis 19.1.4.1  Conventional Techniques Several tests for the identification of streptoccoci belong to the standard laboratory repertoire. These include the Lancefield agglutination test, the GAS quick test and the pneumococcus quick test. Other methods like emm-typing and superantigen determination using PCR are more advanced tests that are usually only performed in more specialized laboratories. Lancefield agglutination test. Streptococci possess specific polysaccharide cell wall antigens, which allow the bacteria to be classified into groups. The antigen is extracted into solution and solubilized antigen is then added to a suspension of latex particles coated with specially purified rabbit antibodies to the group antigens. If a group antigen is present, that group of latex will agglutinate. If no

Western USA United Kingdom

NC, USA

Northern Germany Cambridge, United Kingdom

Jun 1942 Nov 1942

Nov 1943

Sep 1951

MD, USA

NY, USA

CO, USA

AZ, USA

FL, USA

Perm, Russia Ashdod, Israel FL, USA Israel

Feb 1957

Sep 1965

Apr 1968

Jul 1973

Aug 1974

Apr 1975 1976 June 1979 May 1980

Mar 1952

MA, USA

Location

Jul 1941

Date

– Factory Convention Military base

Prison

Community picnic-Indian Reservation

Air Force Academy

University cafeteria

Charity lunch

Army camp

Army base

Army base

Army base Air Force base

Church lunch

Setting

185 447 72 41

290

255

~1200

500– 600 –

43

265

125

~300 89

102

No. of Cases

Sour cream Egg salad Chicken salad Boiled egg salad

Egg salad sandwich

Potato salad (included sliced eggs)a

Tuna salad that contained boiled eggs

Shrimp salad

Egg salad

Custard (the “skin”)

Not identified

Creamed eggs

Not identified Tinned milk

Ham

Food Implicated

One of five inmates who peeled the eggs had a fever and a sore throat; a throat swab specimen was positive for the outbreak strain – – Cook preparing salad had pharyngitis One of the kitchen workers had tonsillitis 3 days before the outbreak; he and five other kitchen workers were positive for the outbreak strain

One of the food handlers had a cough at the time of custard preparation; a throat swab specimen obtained 2 days later had a positive result Food handlers had positive results of testing done 6 days after food preparation Three food handlers who prepared the shrimp salad had negative throat swab specimens Thirty persons present at the salad preparation; one of six of those involved in egg preparation was throat swab positive Four food handlers, one of whom had a son with pharyngitis

One of two cooks had an early stage of scarlet fever – Cook in charge of preparation of tinned milk was infected with streptococci No findings despite thorough investigation 30 cooks; no specific findings

Status of Food Handlers

Table 19.1 Overview of Reported Foodborne Streptococcus Infections in the Past 60 Years

S. pyogenes S. pyogenes Group G S. S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes S. pyogenes

S. pyogenes

Organism Involved

–

– M-nontypeable, T12 Group G streptococcus M3

[32] [33] [34] [35]

[31]

[30]

T3/13/B3264

M9, T9

[29]

[28]

[27]

[26]

[25]

[24]

[22] [23]

[21]

Reference

M-nontypeable, T12

Not typed (coinfection with S. flexneri)

Type 25

Type 9

M5

Griffith 15 Type 9

Griffith 2

Typing of Streptococcus Isolates

262 Molecular Detection of Foodborne Pathogens

OR, USA

NH, USA

TN, USA

Japan MO, USA

Puerto Rico

Venice, Italy

Turkey

Israel

Russia

Israel

May 1981

Jul 1982

Nov 1983

1983 Mar 1984

Aug 1984

Jul 1986

1988

Jun 1988

1989

Apr 1990

Military base

–

Military bases with a central kitchen

–

Five banquets in the same restaurant

Private party

– Conference luncheon

Charity luncheon

Private party

Microbiology conference

61

–

439

58

179

23

– 60

20

34

~300

Boiled egg salad, cabbage salad, and probably egg salad

–

Prawn cocktail in banquet 1, squills and custard cake in banquets 2–4, and no identified food in banquet 5b Bean salad with boiled egg No specific food identified

– Mousse (most probably) and macaroni salad (less likely) Conch salad

Rice dressing

–

No specific food identified

Two of 19 food handlers were symptomatic; both had throat swab specimens positive for the epidemic strain

Two of 12 asymptomatic food handlers had throat swab specimens positive for group A streptococcus (not typed) –

–

Four of ten food handlers had positive throat swabs, and three had skin lesion swabs positive for group A streptococcus A food handler, who was asymptomatic, had a positive throat swab; a household contact had acute pharyngitis shortly before the party Person who had prepared the implicated dish had had pharyngitis 3 weeks earlier and was culture positive at the time of the outbreak – One food handler reported a sore throat; five food handlers had negative throat swabs and no visible hand lesions (8 days later) Four food handlers had negative throat swab specimens and were asymptomatic Six of 18 staff members (five of whom were from the manager’s family) and three other children of the manager were positive for epidemic strain

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

M-nontypeable, T12

–

Probable M29 variant, T-nontypeable

(Continued)

[45]

[44]

[43]

[42]

[41]

M28, T28

M11, T3/13/B3264

[40]

[38] [39]

[34]

[37]

[36]

M-nontypeable, T12, SOR + 

– –

M-nontypeable, T8/25, SOR + 

M-nontypeable, T12, SOR + 

M-nontypeable, T9, SOR + 

Streptococcus 263

LA, USA

Djibouti Israel

–

May 1991

1991 Apr 1992

May 1993 Feb 1991

Aichi Prefecture, Japan Fukuoko Prefecture, Japan Ibaragi Prefecture, Japan New South Wales Australia

Rural correctional ccenter

–

–

Military unit Military field execise Sports meeting

Military base Air Force base

School banquet

– Military base

Church party

Setting

72

–

–

244

162 75

– 197

71

– 75

122, including one death

No. of Cases

Curried egg salad sandwiches

–

Boiled eggs, boiled fish paste, fried chicken, and wakame-gohanc –

Butter Cabbage salad

– White cheese

Macaroni with cheese sauce

– Cabbage salad

Sandwiches, including egg

Food Implicated

b

a

Culture positive for epidemic strain. A chopping board was also found to be culture positive. c All culture positive. Note:  SOR + , serum opacity factor positive. Source:  Modified from Levy, M., Johnson, C.G. and Kraa, E., Clin. Infect. Dis., 36, 175, 2003.

Dec 1999

1997

1997

May 1996

Sweden Israel

1990 Feb 1991

Israel

Sweden

Location

May 1990

Date

Table 19.1  (Continued)

Food handler had infected hand wounds

–

–

Six of seven food handlers had positive throat swab specimens; five of these six attended the party and ate the food. The one who did not attend the party had nasal symptoms 2 days before the party and peeled eggs the day before the party; a second food handler was symptomatic on the day of the party. – Three soldiers were symptomatic before the implicated meal; one was the cook who was throat swab positive for the outbreak strain Food handlers had negative throat swab specimens; one food handler had a positive result of culture of a hand lesion – Food handler was asymptomatic but was throat swab positive; he did not taste the cheese and was absent at the time of the outbreak – Army cook complained of sore throat before suspected meal was served –

Status of Food Handlers

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes

S. pyogenes S. pyogenes

S. pyogenes S. pyogenes

S. pyogenes

S. pyogenes S. pyogenes

S. pyogenes

Organism Involved

–

M56, T25, SOR + 

–

[19]

[56]

[55]

[54]

T1 (PFGE identical)

–

[52] [53]

[50] [51]

[49]

[47] [48]

[46]

Reference

T11 M56, T28

M-nontypeable, T11 M-nontypeable, T 8/25/ Imp19

M9, T5/9

M56, T28

51 of 58 were T28; five of 58 were T28/4; two of 58 were autoagglutinating. The strain were positive for SPEB and SPEC

Typing of Streptococcus Isolates

264 Molecular Detection of Foodborne Pathogens

265

Streptococcus

group antigen is present, the latex stays in homogeneous suspension.58 Kits are commercially available from different companies (Avipath Strep® Omega Diagnostics Ltd.; SlidexTM StreptoKit, bioMerieux). The kits are easy to use and analysis time is in the order of 10–20 min. Most kits identify serogroups A, B, C, D, F, and G. GAS quick tests. GAS quick tests allow for the rapid qualitative detection of GAS antigen directly from throat swab specimens, or from a culture. The tests are lateral flow immunoassays utilizing Quidel’s patented antibody-labeled particles (within 5 min).59 They detect either viable or nonviable organisms directly from throat swabs or culture colonies. The tests are easy to use and are commercially available from different companies (QuickVue Dipstick and QuickVue +   Strep A test, Quidel Corporation; Diaquick, Lab-Stix (Pty) Ltd.; NOW® Strep A Test, Binax). Pneumococcus quick test. The NOW® Streptococcus pneumoniae Antigen Test is an in vitro rapid immunochromatographic assay for the detection of S. pneumoniae antigen in the urine of patients with pneumonia and in the cerebral spinal fluid (CSF) of patients with meningitis. In conjunction with culture and other methods, it is intended to aid in the diagnosis of both pneumococcal pneumonia and pneumococcal meningitis.60 The test is commercially available (NOW® Streptococcus pneumoniae Antigen Test, Binax).

Alternatively, a rapid boiling method can be used: (i) pick half a loop of a fresh overnight culture and resuspend in lysis solution (0.25% w/v SDS, 0.05N NaOH); (ii) boil for 5 min in water bath; (iii) add 900 μl of distilled water; and (iv) ­centrifuge at 11,000 × g for 5 min.

19.1.4.2  Molecular Techniques Molecular techniques for the detection of streptococci are not common. Most methods are designed for subtyping and clonal classification rather than for detection of the bacteria in probe material. Various forms of typing exist including emm-typing and multilocus sequence typing (MLST). Typing of GAS is performed by comparison of the DNA-sequences of the emm-gene, coding for the M-protein, a surface bound virulence factor. S. pneumoniae can be detected in blood and CSF using a PCR amplifying the pneumolysin gene ply. However, since this gene is also sometimes carried by S. mitis and S. oralis strains, the test is not 100% reliable. For most streptococci MLST schemes are available (www.mlst.net). This technique, based on the sequencing of seven housekeeping alleles, is a powerful tool in the detection of clonal relationships among isolates worldwide.

(1) Prepare PCR mixture (100 μl) containing 5 μl of template DNA (as extracted in Section 19.2.1), 10 μl of ten-fold PCR-buffer (500 mM KCl, 100 mM TrisHCl pH 8.3, 15 mM MgCl2, and 0.01% w/v Gelatin), 5 μl of dNTPs (10mM each), 5–20 pmol each of all M forward and reverse primers, 1 μl (5 U) Taq polymerase, and Aqua bidest (to 100 μl). (2) Run the following cycling program: one cycle of 94°C for 2 min; 30 cycles of 94°C for 1 min, 50°C for 1 min, and 72°C for 1.5 min; keep at 4°C until tubes are removed from cycler. (3) Purify the amplified PCR products with standard PCR purification kit (e.g., Qia quick, Qiagen, Germany) and sequence using standard DNA sequencing techniques. (4) Compare the resulting emm-sequences with the emm-database at the Centers for Disease Control (http://www.cdc.gov/ncidod/biotech/strep/strepblast.htm) and assign an emm-type.

19.2  Methods 19.2.1  Sample Collection and Preparation The standard procedures of sampling for streptococci in a foodborne outbreak are food samples, swabs of possibly infected persons and swabs of possibly infected surfaces. Swabs are plated on blood-agar plates, and the colonies are characterized according to their hemolytic characteristics. Isolated strains can be further characterized to determine serogroups using specific antisera. For subtyping using PCR, it is necessary to prepare chromosomal DNA. This is mostly done using commercial DNA-preparation kits. Streptococci can be stored in microbank tubes (CryobankTM, Mast Diagnostica, Germany) at –70°C.

19.2.2  Detection Procedures 19.2.2.1 PCR for emm-Typing Group A Streptococci (GAS) The emm gene encodes the cell surface M-protein. Currently, over 100 different emm-types have been described. PCR with two highly conserved primers allows for the amplification of a large portion of the emm gene. Sequence analysis of the hypervariable sequence encoding M serospecificity allows for the assignment of the different emm-types. The expected size of the PCR fragments is 1–1.85 kb (Figure 19.1). emm types for GAS have the format of ‘emm + a number’ and are usually denoted as: emm1 or emm28. Principle: Podbielski et al.61 described the use of the following primers (all M forward 5’ ATA AGG AGC ATA AAA ATG GCT 3’ and all M reverse 5’ AGC TTA GTT TTC TTC TTT GCG 3’) to amplify the variable part of the emm-gene for typing GAS isolates. Procedure:

Note: an alternative protocol for emm-type determination can be found at http://www.cdc.gov/ncidod/biotech/strep/ protocol_emm-type.htm 19.2.2.2 Multiplex PCR for the Detection of Group A Streptococcal Superantigen In order to detect all 11 SAgs and all allelic variants of SPEA and SMEZ distinctly, a multiplex PCR for GAS SAg has to be performed with two primer sets. In each primer set a 16S rRNA primer for Streptococcus pyogenes is included as a

266

Molecular Detection of Foodborne Pathogens

(a)

Amplificates using ´all M´ primers

emm1 emm4

emm78.3

(b) emm1 GTTTTAGGAG GGGCAACTCA AGAGAAAAAG GATTCTCAGG TCAAAGAACA GCTGAAAAAG GACGCAAGCC CTTGAAGCTA

CAGGCTTTGC CATTACAGCA ATAGATATTT TTGCTGGCCT TGTACGAAAC ATACACTGGC GCAAGAGCCT AACACCAAAA

AAACCAAACA TAAAAATAAT GTATGAAAAA AATAGGTGTA TTTCTTAAAA AGAAAAAGCC AAGCCGTGAC G

GAAGTTAAGG GCATTGACTA GAAGAATTAG GTTGAGAGTG CAGTCTAAAG AAAAAACTTG CTTGAAGCAT

CGGAAGAAGC GTGAGAATGA AAAAGAAAAA ATGAAGAAGA ATCAGGTTAA AAGAAGACAA CTCGTGCAGC

ATCAAATAAT GTCTCTTAGA TAAAGAGTTA AGCTAAGCGC TGAGCTGACT ACAAATCTCA CAAAAAAGAG

60 120 180 240 300 360 420

emm4 GGCTTTGCAA GCGTGGAACT GAACGTGAAA TTAATGGGTG GAACTTGAAA CAAATAGAAG CAATTAGAAG CAAGGTCTAA

ACCAAACAGA GGCCTAAAGA AATATCTATC AAAATCAAGA AAGAAAGAAA CAGATAAGCA CAGAAAAACA GCCGTGACCT

AGTTAAGGCT ATATAACGCG TTATGCTGAC TCTTCGAAAA AGAAAAACAA TTATCAAGAA AAAATTAGCT TGAAGCGTCT

GCGGAGATTA TTACTTAAGG GATAAAGAAA AGAGAGGGAC GAAAGACAAG CAACAAAAGA AAAGACAAAC CGCGAAGCTA

AAAAGCCTCA AAAATGAGGA AAGATCCTCA AATATCAGGA AACAATTAGA AACATCAGCA AAATCTCAGA AGAAAAAAGT

GGCTGATTCA GCTCAAGGTA ATATAGAGCA CAAAATAGAA ACGTCAATAT AGAACAACAA CGCAAGCCGT AGAAGCAGAC

60 120 180 240 300 360 420 480

Figure 19.1  emm-gene PCR. (a) Agarose gel electrophoresis patterns showing PCR amplification products for Streptococcus pyogenes emm-genes. Lanes show (from left to right) PCR products of strains with emm1, emm4 and emm78.3. Although the products are sometimes different in size, they could not be typed by size only and sequencing of the variable region (b) of the amplificate is necessary to define the emm-type. Multiplex 1 RS1 RS6 RS7 SSP4 16SrRNA spea1-3 spel speg spec spek spej

Multiplex 2 RS1 RS6 RS7

16SrRNA ssa spea4 spem smez

Figure 19.2  Superantigen multiplex PCR 1 and 2. Agarose gel electrophoresis patterns showing multiplex PCR 1 amplification product for the Streptococcus pyogenes genes. Left lanes show DNA molecular size marker (50-bp ladder) and the other lanes are indicated by the strain tested. The specific superantigens are labeled with arrows. The figure shows a representative experiment.

positive control. Primer set 1 amplifies 16S rRNA (784 bp), spea alleles 1–3 + 5 (643 bp), spel (460 bp), speg (393 bp), spec (258 bp), spek (233bp), and spej (207 bp). Primer set 2 amplifies 16S rRNA (1042 bp), ssa (570 bp), spea alleles 1–4

(512 bp), spem (411 bp), speh (338 bp), smez (309 bp), and spei (217bp). The two multiplex PCRs show the superantigen profile of the analysed strain and therefore the pathogenicity of the respective strain can be estimated. An example of different superantigen profiles is given in Figure 19.2. The PCR is very reliable. A screening of over 300 isolates resulted in only one SAg negative strain, which was proven to produce no SAg.62 Principle: Lintges et al.62 developed a multiplex PCR with two different primer mixes (Table 19.2) to detect and discriminate all SAg genes. As an internal positive control, each mix contains an additional primer pair targeting the 16S rRNA gene. Procedure: (1) Prepare PCR mixture (50 μl) containing 25 μl master mix (DNA polymerase, dNTP mix, PCR buffer), 5 μl Q-solution (to generate bands of similar intensity), 100 ng template DNA, 200 nM of each SAg gene primer pair, and 20nM 16S rRNA primers. (2) Carry out PCR amplification in a standard PCR thermocycler with an initial denaturation step at

267

Streptococcus

Table 19.2 Composition of Primers used in SAg PCR Primer Mix Mix 1

Target Gene 16S rRNA

f: r: f: r: f: r:

GTGAGTAACGCGTAGGTAACCTACCTCATAG CCCAGGCGGAGTGCTTAATG GGTAAATTTTTCAACGACACACACATTAAA TGTTGAGATTCTCCCGAAATAAATAGAT GCTATGGAAGTCAATTAGCTTATGCAGAT TTATGCGAACAGCCTCAGAGG

f: r: f: r: f:

CAATTAAATTACGCATACGAAATCATACCAGTA ACGAGTAAATATGTACGGAAGACCAAAAATA TATCGCTTGCTCTATACACTACTGAGAGT CCAAACTGTAGTATTTTCATCCGTATTAAA GGACGCAAGTTATTATGGATGCTCA

16S rRNA

r: f: r: f:

TTAAATAAGTCAGCACCTTCCTCTTTCTC GGTATTTGCTCAACAAGACCCCGAT TGTGTTTGAGTCAAGCGTTTCATTATCT GGTGAGTAACGCGTAGGTAACCTACC

r:

GCCCAACTTAATGATGGCAACTAACA

speh

f:

TCTATCTGCACAAGAGGTTTGTGAATGTCCA

r:

GCATGCTATTAAAGTCTCCATTGCCAAAA

spei

f:

AAGGAAAAATAAATGAAGGTCCGCCAT

r: f: r: f: r: f: r: f: r:

TCGCTTAAAGTAATACCTCCATATGAATTCTTT GCTTTAAGGAGGAGGAGGTTGATATTTATGCTCTA CAAAGTGACTTACTTTACTCATATCAATCGTTTC CAATAATTTCTCGTCCTGTGTTTGGAT GATAAGGCGTCATTCCACCATAG AATTATTATCGATTAGTGTTTTTGCAAGTA AGCCTGTCTCGTACGGAGAATTATTGAACTC CAAGAAGTATTTGCTCAACAAGACCCCA TTAGATGGTCCATTAGTATATAGTTGCTTGTTATC

spec speg spej spek spel spea1-3, 5 Mix 2

Primer (5′–3′)

spem smez ssa spea1-4 Note:  f, forward primer; r, reverse primer.

95°C for 15 min, followed by 35 cycles of 94°C for 30 sec, 57°C for 90 sec and 72°C for 90 sec, and a final extension step of 72°C for 10 min. (3) Analyze the samples on a 2.5% agarose gel.

19.3 Conclusion and Future Perspectives Foodborne infections caused by streptococci have become rare in the age of pasteurization and refrigeration. Only about 40 cases of foodborne infections in large groups have been reported over the past 60 years. However, in the developing world, where these preservation methods are not widely available, these infections are most probably still very common. Streptococcal foodborne infections are almost exclusively caused by S. pyogenes (GAS). A foodborne infection with a Group G Streptococcus among attendants of a convention in Florida was reported in 1982.20 The standard method for the analysis of streptococcal foodborne infections is to swab contaminated food and surfaces as well as to take throat swabs of infected patients. Swabs can be cultured on blood agar plates, and a purified culture should be obtained before further analysis with

molecular methods can take place. Streptococci should be grouped using the Lancefield antigen test. GAS should be emm-typed to get insight into the clonal relation between the strains involved in the outbreak. SAg typing of GAS is necessary for further characterization of isolates and assessment of their pathogenicity, and provides, together with the emm-types, an overview of the worldwide strain distribution. Todays high travelling frequency may spread strains from country to country and increase infections due to missing host immunity, which is based on anti-M-proteins and antiSAg antibodies. However, so far GAS has not acquired penicillin resistance, which helps to limit outbreaks.

References

1. Van Bambeke, F. et al. Multidrug-resistant Streptococcus pneumoniae infections: current and future therapeutic options. Drugs, 67, 2355, 2007. 2. Bruckner, D.A., Colonna, P. and Bearson, L.B. Nomenclature for aerobic and faculative bacteria. Clin. Infect. Dis., 29, 713, 1999. 3. Lancefield, R.C. A serological differentiation of human and other groups of hemolytic streptococci. J. Exp. Med., 57, 571, 1933.

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4. Brandt, C.M. et al. Characterization of blood culture isolates of Streptococcus dysgalactiae subsp. equisimilis possessing Lanc­ efield’s group A antigen. J. Clin. Microbiol., 37, 4194, 1999. 5. Reid, S.D. et al. Group A Streptococcus: allelic variation, population genetics, and host-pathogen interactions. J. Clin. Invest., 107, 393, 2001. 6. Hamada, S. et al. Neonatal group B streptococcal disease: ­incidence, presentation, and mortality. J. Matern. Fetal Neonatal Med., 21, 53, 2008. 7. Downar, J. et al. Streptococcal meningitis resulting from contact with an infected horse. J. Clin. Microbiol., 39, 2358, 2001. 8. Sechi, L.A. and Ciani, R. Streptococcus equinus endocarditis in a woman with pulmonary histiocytosis. JAMA, 283, 1005, 2000. 9. Gottschalk, M., Segura, M. and Xu, J. Streptococcus suis infections in humans: the Chinese experience and the situation in North America. Anim. Health Res. Rev., 8, 29, 2007. 10. Mitchell, T.J. The pathogenesis of streptococcal infections: from tooth decay to meningitis. Nat. Rev. Microbiol., 1, 219, 2003. 11. Scott, J.A. The preventable burden of pneumococcal disease in the developing world. Vaccine, 25, 2398, 2007. 12. Bentley, S.D. et al. Genetic analysis of the capsular biosynthetic locus from all 90 pneumococcal serotypes. PLoS Genet., 2, e31, 2006. 13. Pai, R., Gertz, R.E. and Beall, B. Sequential multiplex PCR approach for determining capsular serotypes of Streptococcus pneumoniae isolates. J. Clin. Microbiol., 44, 124, 2006. 14. Butler, J.C. et al. Serotype distribution of Streptococcus pneumoniae infections among preschool children in the United States, 1978–1994: implications for development of a conjugate vaccine. J. Infect. Dis., 171, 885, 1995. 15. Hakenbeck, R. et al. Mosaic genes and mosaic chromosomes: intra- and interspecies genomic variation of Streptococcus pneumoniae. Infect. Immun., 69, 2477, 2001. 16. Hakenbeck, R. et al. Acquisition of five high-Mr penicillinbinding protein variants during transfer of high-level betalactam resistance from Streptococcus mitis to Streptococcus pneumoniae. J. Bacteriol., 180, 1831, 1998. 17. Reinert, R.R. et al. Antimicrobial susceptibility of Streptococcus pneumoniae in eight European countries from 2001 to 2003. Antimicrob. Agents Chemother., 49, 2903, 2005. 18. de Neeling, A.J. et al. Antibiotic use and resistance of Streptococcus pneumoniae in The Netherlands during the period 1994–1999. J. Antimicrob. Chemother., 48, 441, 2001. 19. Levy, M., Johnson, C.G. and Kraa, E. Tonsillopharyngitis caused by foodborne group A streptococcus: a prison-based outbreak. Clin. Infect. Dis., 36, 175, 2003. 20. Stryker, W.S., Fraser, D.W. and Facklam, R.R. Foodborne outbreak of group G streptococcal pharyngitis. Am. J. Epid., 116, 533, 1982. 21. Getting, V.A., Wheeler, S.M. and Foley, G.E. A food-borne Streptococcus outbreak. Am. J. Public Health, 33, 1217, 1943. 22. Bloomfield, A.L. and Rantz, L.A. An outbreak of streptococcic septic sore throat in an army camp. JAMA, 121, 315, 1943. 23. Wilson, M.B. An “explosive” outbreak of haemolytic streptococcal tonsillitis on an RAF station. BMJ, 2, 275, 1944. 24. Commision of Acute Respiratory Disease. A study of food-borne epidemic of tonsillitis and pharyngitis due to b-­haemolytic Streptococcus, type 5. Bull. Johns Hopkins Hosp., 77, 143, 1945.

Molecular Detection of Foodborne Pathogens 25. Gardner, D.L. Streptococcal tonsillitis: an explosive epidemic. J. R. Army Med. Corps, 98, 326, 1952. 26. Boissard, J.M. and Fry, R.M. A food-borne outbreak of infection due to Streptococcus pyogenes. J. Appl. Bacteriol., 18, 478, 1955. 27. Farber, R.E. and Korff, F.A. Foodborne epidemic of group A beta-hemolytic Streptococcus. Public Health Rep., 73, 203, 1958. 28. Elsea, W.R. et al. An epidemic of food-associated pharyngitis and diarrhea. Arch. Environ. Health, 23, 48, 1971. 29. Hill, H.R. et al. Foodborne epidemic of streptococcal pharyngitis at the United States Air Force Academy. N. Engl. J. Med., 280, 917, 1969. 30. McCormick, J.B. et al. Epidemic streptococcal sore throat following a community picnic. JAMA, 236, 1039, 1976. 31. Ryder, R.W. et al. An evaluation of penicillin prophylaxis during an outbreak of foodborne streptococcal pharyngitis. Am. J. Epidemiol., 106, 139, 1977. 32. Lyskovtsev, M.M. et al. Alimentary outbreak of streptococcal angina [in Russian]. Zh. Mikrobiol. Epidemiol. Immunobiol., 134, 1978. 33. Yoshpe-Purer, Y.Y. et al. Food-induced outbreak of streptococcal pharyngitis [in Hebrew]. Harefuah, 91, 248, 1976. 35. Cohen, I.D., Rouach, T.M. and Dinari, G. Food-borne epidemic of streptococcal pharyngitis at an Israeli military training base. Mil. Med., 147, 318, 1982. 36. Dirks, T. et al. A foodborne outbreak of streptococcal pharyngitis—Portland, Oregon. MMWR, 31, 3, 1982. 37. Horan, J.M. and Cournoyer, J.J. Foodborne streptococcal pharyngitis [letter]. Am. J. Public Health, 76, 296, 1986. 34. Decker, M.D. et al. Food-borne streptococcal pharyngitis in a hospital pediatrics clinic. JAMA, 253, 679, 1985. 38. Kashiwagi, Y. et al. A foodborne outbreak of streptococcal sore throat occurring in Tokyo, 1983 [in Japanese]. Kansenshogaku Zasshi, 60, 673, 1986. 39. Martin, T.A. et al. Foodborne streptococcal pharyngitis Kansas City, Missouri. Am. J. Epidemiol., 122, 706, 1985. 40. Berkley, S.F. et al. Foodborne streptococcal pharyngitis after a party. Public Health Rep., 101, 211, 1986. 41. Gallo, G. et al. An outbreak of group A food-borne streptococcal pharyngitis. Eur. J. Epidemiol., 8, 292, 1992. 42. Ulutan, K. et al. A food-borne outbreak of group A streptococcal pharyngitis [in Turkish]. Mikrobiyol. Bul., 23, 302, 1989. 43. Gillis, D. et al. A new Streptococcus group A M-29 variant isolated during a suspected common-source epidemic. Mil. Med., 157, 282, 1992. 44. Sheinkman, E.V. and Kirsanov, I.V. Alimentary outbreak of tonsillitis [in Russian]. Klin. Med. (Mosk), 67, 129, 1989. 45. Lossos, I.S. et al. Food-borne outbreak of group A b-hemolytic streptococcal pharyngitis. Arch. Intern. Med., 152, 853, 1992. 46. Claesson, B.E. et al. A foodborne outbreak of group A streptococcal disease at a birthday party. Scand. J. Infect. Dis., 24, 577, 1992. 47. Niklasson, P.M., Bank, G., and Stattin, U. Utbrott av livsmedelsburen streptokocktonsillit [in Swedish]. Epid. Akt., 8, 2, 1990. 48. Shemesh, E. et al. An outbreak of foodborne streptococcal throat infection. Isr. J. Med. Sci., 30, 275, 1994. 49. Farley, T.A. et al. Direct inoculation of food as the cause of an outbreak of group A streptococcal pharyngitis. J. Infect. Dis., 167, 1232, 1993. 50. Bercion, R. et al. Clinical and biological aspects of a collective alimentary toxi-infection by group A Streptococcus in a military unit stationed in Djibouti [in French]. Bull. Soc. Pathol. Exot., 86, 29, 1993.

Streptococcus 51. Bar Dayan, Y. et al. Food-borne outbreak of streptococcal pharyngitis in an Israeli Airforce base. Scand. J. Infect. Dis., 28, 563, 1996. 52. Briko, N.I. et al. An explosive outbreak of strep throat ­morbidity in an adult organized collective [in Russian]. Zh. Mikrobiol. Epidemiol. Immunobiol., September/October, 35. 1993. 54. Matsumoto, M. et al. An outbreak of pharyngitis caused by food-borne group A Streptococcus. Jpn. J. Infect. Dis., 52, 127, 1999. 55. National Institute of Infectious Diseases. Outbreak of group A streptococcal infections caused by contaminated luncheons— Fukuoka Prefecture [in Japanese]. Infect. Agents Surveill. Rep., 18, 264, 1997. 56. National Institute of Infectious Diseases. Outbreak of group A streptococcal infections caused by contaminated luncheons provided by a caterer in Ibaragi Prefecture (in Japanese). Infect. Agents Surveill. Rep., 19, 279, 1997. 53. Shemesh, E. et al. An outbreak of foodborne streptococcal throat infection. Isr. J. Med. Sci., 30, 275, 1994. 57. Wannamaker, L.W. Perplexity and precision in the diagnosis of streptococcal pharyngitis. Am. J. Dis. Child., 124, 352, 1972.

269 58. Petts, D.N. Evaluation of a modified nitrous acid extraction latex agglutination kit for grouping beta-hemolytic streptococci and enterococci. J. Clin. Microbiol., 33, 1016, 1995. 59. Nerbrand, C., Jasir, A. and Schalén, C. Are current rapid detection tests for Group A Streptococci sensitive enough? Evaluation of 2 commercial kits. Scand. J. Infect. Dis., 34, 797, 2002. 60. Petti, C.A., Woods, C.W. and Reller, L.B. Streptococcus pneumoniae antigen test using positive blood culture bottles as an alternative method to diagnose pneumococcal bacteremia. J. Clin. Microbiol., 43, 2510, 2005. 61. Podbielski, A., Melzer, B. and Lütticken, R. Application of the polymerase chain reaction to study the M protein(-like) gene family in beta-hemolytic streptococci. Med. Microbiol. Immunol., 180, 213, 1991. 62. Lintges, M. et al. A new closed-tube multiplex real-time PCR to detect eleven superantigens of Streptococcus pyogenes identifies a strain without superantigen activity. Int. J. Med. Microbiol., 297, 471, 2007.

Section III Foodborne Gram-Negative Bacteria

20 Aeromonas

Germán Naharro, Jorge Riaño, Laura de Castro, Sonia Alvarez, and José María Luengo University of León

Contents 20.1 Introduction.................................................................................................................................................................... 273 20.1.1 General Consideration ..................................................................................................................................... 273 20.1.2 Aeromonas Biology and Taxonomy.................................................................................................................. 274 20.1.3 Aeromonas Epidemiology................................................................................................................................. 276 20.1.4 Aeromonas Virulence....................................................................................................................................... 276 20.1.4.1 Cell-Associated Virulence Factors.................................................................................................. 277 20.1.4.2 Extracellular Virulence Factors....................................................................................................... 278 20.1.5 Isolation and Biochemical Identification ......................................................................................................... 278 20.1.5.1 Isolation............................................................................................................................................ 278 20.1.5.2 Biochemical Identification............................................................................................................... 279 20.1.6 Molecular Detection and Identification............................................................................................................ 279 20.2 Methods.......................................................................................................................................................................... 281 20.2.1 Reagents and Equipment................................................................................................................................... 281 20.2.2 Sample Collection and Preparation.................................................................................................................. 281 20.2.3 Detection Procedures........................................................................................................................................ 281 20.3 Conclusions and Future Perspectives............................................................................................................................. 282 References.................................................................................................................................................................................. 283

20.1  Introduction 20.1.1  General Consideration The genus Aeromonas is a member of the family Aeromonadaceae and is an inhabitant of aquatic environments. Some species have been described as pathogens for humans and animals1 and one of them, A. hydrophila has been included in the First and Second Contaminant Candidate List (CCL1 and CCL2) by the Environmental Protection Agency (EPA) of the USA of potential waterborne pathogens.2,3 Further, the EPA Method 16054 has been validated for the detection and enumeration of A. hydrophila in drinking water. Aeromonas species are widely distributed in nature, occurring not only in aquatic environments—including ground water, surface water, estuarine and marine water, drinking water, and wastewater5—but also in different fresh and raw foods,6 which is why they are considered as potential agents in foodborne diseases. Aeromonas species may cause several types of infection in man such as septicaemia, usually associated with immunosuppression resulting from hepatitis, cirrhosis, biliary disease, pancreatic disease or malignancies; wound infections and cellulite in individuals exposed to water or soil; extraintestinal infections such as peritonitis, meningitis, endocarditis, otitis of the middle ear, etc.7 Several reports have implicated Aeromonas species as the etiological agents of acute diarrhea, but the

evidence of enteropathogenicity is controversial because the ­experimentation undertaken to solve the issue has proven insufficient, and conclusive human volunteer trials and animal models are required.8,9 Aeromonas is not considered to be a normal inhabitant of the human gastrointestinal tract, but it is present in the feces of healthy animals and humans, probably due to the ingestion of water and foods contaminated with these microorganisms.10,11 Aeromonas may form a biofilm after the colonization of drinking water distribution systems, where it is resistant to common disinfectants. However, no outbreaks attributable to Aeromonas through contact with drinking water have been described.12 The taxonomy of the genus Aeromonas is complex and confusing because of a lack of congruity between the phenotypic and genotypic properties of the various species based on biochemical characteristics and DNA-DNA hybridization characteristics, respectively, and multiple methods are required for accurate classification, which exceeds the analytical capacity of most clinical laboratories. In the most recent edition of Bergey’s Manual, Martin-Carnahan and Joseph1 describe 17 DNA–DNA hybridization groups (HG) and 14 phenospecies are reported. Several Aeromonas species have recently been added to the list, including A. simiae, A. molluscorum, A. culicicola, although the latter may be synonymous of A. veronii. Some species are synonymous of previously recognized species, such as A. ichthiosmia 273

274

(A. veronii biovar sobria) and A. enteropelogenes (A. trota). Some researchers consider the continued use of HG to be controversial, since all phenospecies named can be identified phenotypically. An Aeromonas infection in humans was first reported in 1954 in a woman from Jamaica with myositis.13 In 1968, von Graevenitz and Mensch reviewed 30 cases of infections and established Aeromonas species as significant human pathogens, causing a variety of extraintestinal infections in various organ systems.14 Aeromonas infections typically follow trauma in an aquatic environment, or dissemination from the gastrointestinal tract as a result of underlying diseases such as cancer. Human infections are known to occur in both healthy hosts and immunocompromised individuals.15–17 Aeromonas infections in humans are mainly produced by A. hydrophila (HG1), A. caviae (HG4), A. veronii biovar sobria (HG8), A. jandei (HG9), A. veronii biovar veronii (HG10), A. schuvertii (HG12), and A. trotta (HG14). Probably, A. trotta (HG14) has been underestimated, since it is sensitive to ampicillin, an antibiotic included in most Aeromonas isolation media.17 Fortunately, 85% of human clinical isolates are A. hydrophila (HG1), A. caviae (HG4), and A. veronii biovar sobria (HG8). Aeromonas species produce virulence factors, including cell-associated virulence factors and extracellular virulence factors. Among the cell-associated virulence factors are surface structures such as pili, flagella, outer-membrane proteins, lipopolysaccharide, and capsules. The major extracellular products are cytotonic, cytolytic, hemolytic, enterotoxic, lipolytic, and proteolytic proteins.16,18,19 Some Aeromonas species have been found to be pathogenic for fish, amphibians, and reptiles. In fish they are the etiological agents of hemorrhagic disease, ulcerative disease, furunculosis, and septicemia.20,21 Aeromonas species have also been described as pathogenic for birds and domestic animals, causing abortion, pneumonia, peritonitis, and various other diseases.21,22

20.1.2  Aeromonas Biology and Taxonomy Aeromonas species are natural inhabitants of aquatic environments, where they may be primary pathogens for poikylotherms (cool-blooded animals). They are found in soils, sediments, and water in freshwater, estuarine, and marine environments. They are present in foods, sewage, and drinking water distribution systems. Aeromonas are found in the intestines of animals and humans where they may or may not cause gastrointestinal disease. Some aeromonads are pathogenic for humans, and most human clinical isolates belong to HG1, HG4, HG8, HG9, HG10, HG12, or HG14.17 The proportion of strains within these HG able to cause human disease is unknown. HG2, HG3, HG5, HG6, HG7, HG11, HG15, HG16, or HG17 are isolated from the environment or for diseased animals, and they are not usually considered to be human pathogens. Some strains of Aeromonas produce enterotoxins responsible for gastroenteritis in humans; however, the isolation of aeromonads from feces does not prove pathogenicity,

Molecular Detection of Foodborne Pathogens

since these bacteria are widely distributed in water and foods, especially in the summer.17 The genus Aeromonas consists of straight coccobacillary to bacillary Gram-negative bacteria with round ends measuring 0.3–1.0 × 1.0–3.5 µm.1 They occur singly, in pairs, and— although rarely—in short chains. Motile strains produce a single polar flagellum, but peritrichous or lateral flagella may be formed by some species in solid media. Aeromonas spp. are facultatively anaerobic, catalase-positive, oxidase-positive, chemoorganotrophic bacteria with a fermentative and oxidative carbohydrate metabolism. Aeromonas spp. produce a broad variety of extracellular hydrolytic enzymes such as arylamidases, amylase, deoxyribonuclease, esterases, peptidases, elastase, chitinase, and lipase.23–25 Aeromonas spp. grow optimally within a temperature range between 22 and 35ºC, although some species are able to grow in the 0–45ºC temperature range.26 Some species, including most nonmotile A. salmonicida strains, do not grow at 35ºC.1 Aeromonas spp. tolerate a pH range from 4.5 to 9.0, but the optimum pH range is from 5.5 to 9.0,27 and the optimum sodium chloride concentration range is from 0 to 4%. Aeromonas lipopolysaccharide, the major surface antigen (O) as in other Gram-negative bacteria, is the main parameter used in serotyping.28 Several typing schema have been proposed29–31 but few comparisons have been made in the literature.32 The scheme proposed by Sakazaki and Shimada28 recognizes 44 serogroups, with an additional 52 provisional serogroups proposed by others.33 Aeromonas spp. have been found to be serologically heterogeneous, with individual serogroups encountered in more than one species.34 Most type and reference strains are not serologically representative of a genomospecies. Polyclonal antibodies for the rapid identification of clinical isolates by direct agglutination of A. hydrophila have been developed, and the distribution of serogroups in clinical specimens has been studied.35 In that study only rough strains (15.2%) and untypable strains (2.3%) reduced the effectiveness of serotyping for the identification of clinical strains. Most motile aeromonads are generally resistant to penicillin, ampicillin, carbenicillin, and ticarcillin, and they are typically susceptible to second- and third-generation cephalosporines, aminoglycosides, carbapenems, chloramphenicol, tetracyclines, trimethoprim-sulfamethoxazole, and quinolones.17,36 Most aeromonads produce inducible chromosomal β-lactamase37,38 but A. trota and up to 30% of A. caviae are susceptible to ampicillin. Antibiotic resistance to streptomycin, chloramphenicol, tetracycline, cephalexin, erythromycin, furazolidone, and sulfathiazole is mediated by plasmids.39 The taxonomy of Aeromonas has been extensively revised during the last 20 years and became complicated in the 1980s because few phenotypic markers were available to differentiate species for the newly recognized HG. The first observation of Aeromonas is attributed to Zimmermann in 1890,40 who considered these microorganisms as belonging to the genus Bacillus (Bacillus punctatus). In 1891, Sanarelli41 isolated a strain from infected frog blood: it was named Bacillus

275

Aeromonas

hydrophilus fuscus and was considered to be an aeromonad. In the middle of the twentieth century the nomenclature of the genus Aeromonas was still not clear; both Stanier42 and Miles43 suggested that many strains included in several genera, such as Pseudomonas, Proteus, Bacillus, and Acetobacter, in fact belonged to the genus Aeromonas. In 1936, Kluyver and van Neil44 coined the name Aeromonas, from the Greek words aer (air or gas) and monas (unities) which means “gas producing units.” The genus Aeromonas would include Gram-negative, facultatively anaerobic, rodshaped bacteria (1–3.5 µm), although cocci may also appear, with diameters ranging from 0.3 to 1.0 µm. The microorganisms are chemoorganotrophic, with an optimum growth temperature of 30ºC, a minimum growth temperature of 4ºC and a maximum growth temperature of 45ºC. Oxidase- and catalase-positives, they reduce nitrate, ferment D-glucose, and are resistant to the vibriostatic agent O129. The natural habitat of Aeromonas is in watery environments associated with aquatic animals and sediments. The genus Aeromonas was included in the family Pseudomonadaceae in the 1957 seventh edition of Bergey’s Manual45 with three motile species (A. hydrophila, A. punctata, and A. liquefaciens) and one nonmotile species (A. salmonicida). Schubert, in the 1974 eighth edition of Bergey’s Manual, moved the genus Aeromonas to the family Vibrionaceae and, based on their phenotypic properties, divided motile aeromonads into two species: A. hydrophila, with three subspecies (A. hydrophila hydrophila, A. hydrophila anaerogenes, and A. hydrophila proteolitica) and A. punctate, with two subspecies (A. punctata punctata and A. punctata caviae), whereas the nonmotile species A. salmonicida included three subspecies (A. salmonicida salmonicida, A. salmonicida achromogenes, and A. salmonicida masoucida).46 In 1976, Popoff and Veron47 carried out an interesting study based on numeric taxonomy in which they differentiated two taxonomic groups among the motile species: the A. hydrophila group, with two biovars (A. hydrophila biovar hydrophila and A. hydrophila biovar anaerogenes), and the A. sobria group, corresponding to a new species. However, further studies using DNA–DNA hybridization techniques demonstrated that A. hydrophila biovar anaerogenes was a member of the species A. caviae. Further, the species A. hydrophila, A. sobria, and A. caviae included two or three HG each that could not be identified by phenotypic methods.48 All these data are recorded in the 1984 first edition of Bergey’s Manual of Systematic Bacteriology,49 considering three mesophilic motile species: A. hydrophila, A. sobria, and A. caviae, and one psychrophile nonmotile species: A. salmonicida, with three subspecies: A. salmonicida salmonicida, A. salmonicida achromogenes, and A. salmonicida masoucida. Each of these four species has two or more HG: three in A. hydrophila (HG1, HG2, HG3), two in A. caviae (HG4, HG5; although HG5 was later associated with A. media), and at least two in A. sobria (HG7, HG8). Aeromonas taxonomy was further complicated in the 1980s because different researchers published genetically identical strains under different names. DNA–DNA HG

were recognized for which there were no reliable phenotypic ­characteristics, resulting in confusion among microbiologists and physicians. Newly recognized species of Aeromonas were described from just a few isolates, making it difficult to describe the species phenotypically. This was the case of A. allosaccharophila, which was described from only three isolates;50 A. encheleia was described from four isolates,51 and A. popoffii was described from just eight isolates.52 According to the second edition of Bergey’s Manual of Systematic Bacteriology1,53 the genus Aeromonas includes 14 phenospecies and at least 17 HG, although the list is not closed since new species are continuously added (Table 20.1). The genus Aeromonas belongs to the family Aeromonadaceae, order Aeromonadales, subclass γ-Proteobacteria, class Proteobacteria. Current taxonomy is based on DNA–DNA hybridization analysis and 16S rRNA sequences. Nevertheless, some discrepancies have been observed between 16S rRNA sequences and DNA–DNA hybridization results owing to the high degree of conservation of 16S rRNA sequences (97.8– 100%)54 and the existence of several copies of the gene with intragenomic gene polymorphism, especially in A. media and A. veronii.55 Recently, some investigators have started to use sequences other than 16S rRNA to identify Aeromonas to the genomic species level, such as rpoB,56 gyrB,57 dnaJ,58 and recA.59 This multigene approach to these housekeeping genes provides better molecular markers than 16S rRNA for the study of phylogenetic and taxonomic relationships at species level and fulfills the recommendations of the Table 20.1 Current Genomospecies and Phenospecies of the Genus Aeromonas HG

Type Strain

Genomospecie

1 2 3

ATCC 7966 ATCC 14715 ATCC 33658

A. hydrophila A. bestiarum A. salmonicida

4 5A 5B 6 7 8X 8Y 9 10

ATCC 15468 CDC 0862-83 CDC 0435-84 ATCC 23309 CIP 7433 CDC 0437-84 ATCC 9071 ATCC 49568 ATCC 35624

11

ATCC 35941

A. caviae A. media A. media A. eucrenophila A. sobria A. veronii A. veronii A. jandei A. veronii biovar veronii Unnamed

12 13

ATCC 35941 ATCC 43946

14 15 16 17

ATCC 49657 CECT 4199 CECT 4342 LMG 1754

A. schubertii Aeromonas Group 501 A. trota A. allosaccharophila A. encheleia A. popoffii

Phenospecie A. hydrophila A. hydrophila-like A. salmonicida ssp salmonicida A. caviae A caviae-like A. media A. eucrenophila A. sobria A. sobria A. veronii biovar sobria A. jandei A. veronii biovar veronii Aeromonas spp. (ornithina positive) A. schubertii A. schubertii-like A. trota A. allosaccharophila A. encheleia A. popoffii

276

ad hoc committee for the re-evaluation of the definition of the ­bacterial species.60

20.1.3  Aeromonas Epidemiology Some Aeromonas species are opportunistic pathogens of human beings, causing a variety of extra-intestinal infections as well as gastrointestinal disease. von Graevenitz and Mensch published the first comprehensive review of human infections caused by Aeromonas species.14 Since then, a vast number of reviews and papers has been published on Aeromonas species as agents of human disease,7,15,16,61,62 such as cellulite, abscesses, wound infection, necrotizing fasciitis, myonecrosis, pneumonia, empyema, septicemia, septic arthritis, osteomyelitis, endocarditis, meningitis, gastroenteritis, appendicitis, peritonitis, acute suppurative cholangitis, and corneal ulcer. The clinical symptoms of Aeromonas infections depend on the site and severity of infection. Septicemia may follow wound infection, and dissemination may result in meningitis or endocarditis. Patients presenting gastroenteritis symptoms range from self-limiting diarrhea to acute, severe diarrhea with abdominal cramp, vomiting, and fever. Children under 12 are likely to have more acute and severe illnesses, whereas adults may have chronic diarrhea and abdominal cramps. A. caviae and A. hydrophila are more associated with chronic diarrhea.63 Aeromonas species are ubiquitous and find multiple opportunities for transmission to humans through food, water, animal contact, and direct human contact. Extraintestinal infections may be acquired after trauma in aquatic environments, and intestinal infections are acquired by ingestion of contaminated food or water. Additionally, intestinal infections in immunocompromised patients may disseminate, resulting in septicemia with multiple organ involvement. Aeromonas species have been recognized as significant agents in foodborne illnesses, gastroenteritis being the most commonly reported infection that range from mild, selflimiting watery diarrhea to a more severe, invasive Shigellalike dysenteric form.62,64 Aeromonas is frequently isolated from foods, such as green vegetables, raw milk, ice cream, meat, and seafood.65–67 von Graevenitz defined Aeromonasassociated diarrhea as loose stools associated with the isolation of aeromonads in the stools.62 However, the evidence of outbreaks with aeromonads is controversial and almost nonexistent in spite of their occurrence in drinking water68 and in foods such as seafood, meat, and vegetables.6 The problem arises because of differences in the diarrheagenic potential among Aeromonas strains. A general consensus appears to be growing that certain strains are likely to be human enteric pathogens.17,69 Several typing methods have been used to link environmental and clinical strains of aeromonads; however, the diversity of strains has resulted in little success in all but a few cases. Ribotyping was used to demonstrate that the ingestion of shrimp resulted in gastroenteritis in the first report of foodborne illness attributed to Aeromonas species.70 The same method was used to demonstrate that a patient with chronic diarrhea carried the same strain for years71 and to

Molecular Detection of Foodborne Pathogens

demonstrate person-to-person transmission of Aeromonas species between a foster child and a foster parent.72 Only four phenospecies of the genus Aeromonas have been isolated from human feces with significant frequency: A. hydrophila (HG1), A. caviae (HG4), A. veronii biovar sobria, (HG8), and A. trota (HG14).1,7 A. jandei (HG9) and A. schubertii (HG12) have been rare fecal isolates.73 In any case, it is to be expected that if these species are indeed agents of diarrhea, they should be more frequently isolated from patients with Aeromonas-associated diarrhea than in controls. Several studies conducted to address this issue have demonstrated that the epidemiological behavior of Aeromonas is different from that of other enteropathogenic bacteria, and there were even differences in the geographical and seasonal distributions, and the predominant species involved. Some studies observed isolation peaks in warmer seasons,74–76 while others found no seasonal preferences.77 A. caviae was the predominant species in most studies, followed by A. hydrophila and A. veronii biovar sobria.8,75–78 A. hydrophila predominates in Brazil and Thailand.79,80 A. caviae initially predominated in Bangladesh, but later A. trota was held responsible.81,82 In Finnish tourists visiting Morocco, diarrhea was attributed to A. veronii biovar sobria.83 While some studies did find significant differences in the frequency of aeromonads isolated from diarrheic patients including travellers83 as compared to controls,74,76,77 others did not.78,84,85 The recovery rates of aeromonads from children with diarrhea vary geographically: 0.62–4% in Malaysia,86 0.75% in Nigeria,87 2% in Sweden,85 4.8% in Switzerland,88 and 6.8% in Greece.89 In Thailand, the difference was significant for western peace corps workers but not for indigenous people.80 In a quantitative study, a considerable variation in the number of colony-forming units (CFU) of aeromonads during different sampling periods was observed between patients and controls.90 In another study the amount of growth on selective medium was not found to be associated with the onset of diarrhea8.The isolation rate for human fecal specimens varies widely, since the geographical area, patient population, food habits, level of sanitation, and culture methods influence the recovery rate.91

20.1.4  Aeromonas Virulence The virulence of a pathogen will depend of four basic capacities: adherence to the mucous surfaces of the host and colonization; entry into host tissues, growth in host tissues, and resistance to host defence mechanisms and host injury. The virulence of aeromonads is multifactorial and incompletely understood despite years of intensive investigation. Many putative virulence factors have been described, and have usually been grouped into two classes: cell-associated virulence factors (outer-membrane proteins, S or A layer, lipopolysaccharide, capsule, flagella, and pili), and extracellular virulence factors (toxins, enterotoxins, proteases, hemolysins, and lipases, etc.).92,93 In the early literature only one virulence factor—the A-layer of A. salmonicida—was unequivocally linked to disease causation. However, much progress

Aeromonas

has been made in understanding the virulence mechanisms of these microorganisms since then.94 20.1.4.1  Cell-Associated Virulence Factors Invasins. Intracellular Aeromonas spp. have been demonstrated by electron microscopy, although no gene or gene product has been identified specifically with invasion, their invasive ability proving to be similar to that of Campylobacter.95 Adhesins. Adherence ability at the Aeromonas surface96 has been reported, together with a correlation between highlevel HEp-2 cell adherence and enteropathogenicity.97 Outer-membrane proteins. It is generally accepted that outer membrane proteins (OMP) are involved in the adherence of Aeromonas spp. to cells. A 43-kDa OMP from A. caviae promoting adhesion to Hep-2 cells has been reported.98 This 43-kDa OMP expressed higher levels when A. caviae was grown at 22ºC than when grown at 37ºC. Similar adherence characteristics have been observed in A. veronii biovar sobria 99 and A. hydrophila,100 which have been characterized at molecular level. S-layer proteins. The S-layer (also termed A-layer) has been extensively studied and has been linked to virulence in that it provides protection against bactericidal serum ability, protecting cells from proteolysis, facilitating association with macrophages, binding collagen, laminin, fibronectin, and playing a role in colonization.101 The S-layer, which surrounds the outer membrane of aeromonads and indeed many other Gram-negative, Grampositive bacteria and archaea, exhibits a paracrystalline array made up of protein units with a molecular mass of 49–51 kDa. The S-layer-coding genes from A. salmonicida (vapA) and A. hydrophila (ahs) have been fully characterized.102 When these genes are insertionally inactivated, the S-layer is lost and virulence is markedly reduced.102 S-layers are thought to protect against bacterial parasites such as Bdellovibrio bacteriovorus, although they are ineffective against protozoa.102 Lipopolysaccharide. Lipopolysaccharide is one of major surface antigens (O-antigen) of the Gram-negative bacterial cell wall that consist of three domains: Lipid A, core oligosaccharide, and the distalmost O-specific polysaccharide or O antigen. Lipopolysaccharide plays a role in adhesion to epithelial cells,103 resistance to nonimmune serum,104 and virulence.105 The serotyping of aeromonads is based on the distalmost O-antigen of LPS. LPS is thought to be a colonization factor since its presence facilitates adherence to HEp-2 cells. The loss of LPS reduces adhesion by 60%.106 Many studies have been performed on the structure, function, genetic characterization and role in virulence of the O-polysaccharide of A. hydrophila O:34.107–109 Capsule. Most motile aeromonads have no capsule, so few studies have examined the role of capsule polysaccharide as a virulence factor. Capsules have been reported in A. hydrophila serotypes O:11 and O:34 when grown in a glucose-rich medium.50 The capsule may play a role in septicemia since non-encapsulated strains are less virulent. The capsule material has the ability of to protect cells from ­complement-mediated serum killing activity.110

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Flagella. Aeromonas generally have monotrichous and polar flagella. However, some strains present lateral flagella, and some strains are nonflagellated. Peritrichous flagella are unsheathed and are associated with a swarming movement across the surfaces of solid media.96 Flagellar production is under the control of over 40 genes and allows Aeromonas to reach target cells, where they colonize.111 Lateral flagella facilitate biofilm formation and persistence during infection.112 Polar sheathed flagella are expressed constitutively in liquid media and peritrichous lateral flagella are produced in matrices that do not permit motility by a single polar flagellum, such as growth on solid media.113,114 The expression of lateral flagella and their multifunctional role in pathogenesis has been reviewed, and the flagellar genes have been identified.115 Lateral flagella permit swarming motility on surfaces and function as adhesions, contributing to microcolony formation and biofilm formation on surfaces. Both lateral and polar flagella are enterocyte adhesins that contribute to biofilm formation on surfaces.116 About 60% of mesophilic aeromonads produce lateral flagella. Strains that lose polar flagella are virtually nonadherent to cell lines, while strains lacking lateral flagella have a reduced capacity of cell adherence. Eighty percentage of binding capacity is lost in flagellar mutants. Pili (fimbriae). Aeromonas attachment to host cells is mediated by surface appendages such as pili which have been described as potential colonization factors in A. hydrophila and A. veronii biovar sobria.117 Two morphotypes of pilus have been observed and reported to be important adhesive factors for mucosal surface attachment in Aeromonas species: short rigid pili (S/R type) and long wavy flexible pili (L/W type).115 The S/R type of pilus facilitates self-pelleting but not hemagglutination and is present in 95% of strains, regardless of their source,99 although S/R pili have also been described to be more abundant in environmental strains.118 S/R pili have been purified from A. hydrophila; they have a diameter of 7–10 nm, and the major protein—pillin—has a molecular mass of 17 kDa, showing homology in 28 of 40 residues with E. coli Type I and the Pap-pyelonephritisassociated pili.118 L/W type pili cause hemagglutination, are associated with OMP, and are found on clinical isolates.119 This type of pilus has a diameter of 4–7 nm and can form bundles or filamentous networks, showing maximum expression when grown at 22ºC.99,119 Removal of these pili blocks adherence and when they react with cell receptors the competitively block the adherence of the parental bacterial strains.120 The main component of Aeromonas pili is pilin which share sequence similarity with Type IV pilins of other Gram-negative bacteria although they are immunologically and electrophoretically distinct. The Type IV pilins are further subdivided into two groups—Type IVa and Type IVb—on the basis of the lengths of their signal peptide and mature sequence.121 Both types of pilus have been described in Aeromonas species from cases of gastroenteritis;122,123 however, the contribution of these pili to virulence needs further studies.124

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20.1.4.2  Extracellular Virulence Factors Proteases. Most aeromonads produce a broad variety of extracellular enzymes, some of which may be involved in pathogenesis.125 Among them, proteases are thought to play a significant role in aeromonad virulence. It has been described that aeromonads differ in the elaboration of proteases according to differences in strain origin, incubation temperature, and the culture medium.26 Most strains of A. hydrophila and A. veronii biovar sobria produce two types of proteases, a thermostable metalloprotease (TSMP) that is sensitive to EDTA, and a thermolabile serine protease (TLSP) that is sensitive to PMSF.126 The two major extracellular proteolytic activities of A. hydrophila described in our lab, a 38-kDa TSMP24 and a 68-kDa TLSP,25 are present in most A. hydrophila culture supernatants. We have also demonstrated that the 38-kDa TSMP, with high elastolytic activity, is essential for virulence in rainbow trout while the 68-kDa TLSP is not.18,19 Lipases. Several lipase-encoding genes have been cloned for Aeromonas, and in some cases mutation of these genes reduces lethal effects in mice and fish;127 however, we have constructed an A. hydrophila lipase mutant by allelic replacement and the lethal effects in fish were not reduced (unpublished results). Glycerophospholipid cholesterol acyl-transferase (GCAT). GCAT activity has been described as a major virulence factor in A. salmonicida resulting in furunculosis in fish. A similar enzyme has also been described in A. hydrophila that hydrolyzes phospholipids. However the role of GCAT in human pathogenicity is unknown.128 Superoxide dismutase. An A. hydrophila manganese superoxide dismutase has been described with an equivalent function to the copper/zinc superoxide dismutase of E. coli. This enzyme is located in the periplasmic space of A. hydrophila and may confer a virulence advantage on the bacteria.129 Toxins. Aeromonas species produce several toxins considered as virulence factors, such as hemolysins, and cytotoxic and cytotonic enterotoxins. The first toxin purified to homogeneity from an Aeromonas was a heat-labile 49- to 52-kDa β-hemolysin, that induced fluid secretion in an animal model.130 This is the major hemolysin produced by Aeromonas and is called aerolysin (AerA), although it is also known by other names (cytotoxin enterotoxin, Asao toxin, and cholera toxin cross-reactive cytolytic enterotoxin). Aerolysin is produced by some strains of A. hydrophila, A. veronii biovar sobria, and A. caviae.131 The aerolysin coding gene, aerA, is present in all Aeromonas strains but its expression depends on the prevailing environmental conditions.131 Aerolysin is a known virulence factor synthesized as an inactive precursor, with a 25-amino acid leader peptide and a 25-amino acid C-terminus that must be removed for activation. Aerolysin belongs to a class of pore-forming cytotoxins that disrupt cell membranes and are highly lethal for rats and mice. In vitro they cause fluid accumulation in rabbit ileal loops and lyse a wide range of cells, including CHO and rabbit erythrocytes.132

Molecular Detection of Foodborne Pathogens

Furthermore, three cytotoxic and cytotonic ­enterotoxins have been described in Aeromonas species: Aeromonas cytotoxic enterotoxin (Act), Aeromonas heat-labile (56ºC) cytotonic enterotoxin (Alt), and Aeromonas heat-stable cytotonic enterotoxin (Ast).133 Act is a single-chain polypeptide of 52-kDa that is related to aerolysin and has hemolytic, cytotoxic, and enterotoxic activities. The role of Act in Aeromonas virulence has been clearly demonstrated by LD50 studies and by the inability of act isogenic mutants to elicit a fluid secretory response in mouse ligated ileal loops.134 Alt is a heat-labile cytotonic enterotoxin exhibiting a size of 44-kDa single polypeptide that causes elevation of cyclic AMP and prostaglandin levels in CHO and intestinal epithelial cells, resulting in fluid accumulation in rat ileal loops, while Ast is a heat-stable cytotonic enterotoxin with a similar molecular mass and properties.

20.1.5  Isolation and Biochemical Identification 20.1.5.1  Isolation A variety of selective and differential media have been proposed to isolate Aeromonas species, and even broth enrichment methods are frequently used to recover aeromonads from heavily contaminated samples.135 There are more than 17 genomospecies of Aeromonas; however, only seven associated with human diarrhea and include A. hydrophila, A. caviae, A. veronii biovar sobria, A. veronii biovar veronii, A. schubertii, A. jandaei, and A. trota. Eighty-five percent of clinical isolates belong to A. hydrophila (HG1), A. caviae (HG4), and A. veronii biovar sobria (HG8).136 Selectivity is based on the use of media supplemented with ampicillin and/or inhibitors used in culture media for Enterobacteriaceae (bile salts, brilliant green, and sodium lauryl sulphate). Differential media are most often based on the presence of amylase in the microorganisms to be isolated and the pattern of carbohydrate fermentation, but no single medium has yet received general acceptance. Several studies have been performed by comparing a series of selective media with or without enrichment in alkaline peptone water (APW). Some authors have concluded that ampicillin dextrin agar (ADA) produces the best overall results; however, this selective medium is not appropriate for the isolation of A. trota and certain strains of A. caviae, because they are sensible to ampicillin.137–139 ADA medium supplemented with vancomycin (ADA-V) has been validated by the USEPA for the detection and enumeration of A. hydrophila in drinking water.4 Starch-ampicillin agar, Aeromonas medium, and starch glutamate-ampicillinpenicillin media were used in the quantitative recovery of Aeromonas species from foods. It was found that in heavily contaminated foods, the Aeromonas medium was the most adequate one for the recovery of Aeromonas species, especially A. sobria, and in moderately contaminated foods any of the three media may be used for the recovery of A. hydrophila and A. caviae.140 One interesting study evaluated seven different selective media and two enrichment

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Aeromonas

broths for the isolation of mesophilic Aeromonas from meat and fish. It was concluded that ­enrichment in APW, and consecutive plating in two selective media—ampicillin-sheepblood agar supplemented with 30 µg/ml of ampicillin (ASBA 30) and bile salts-Irgasan-brilliant green agar (BIBG) should be recommended for qualitative isolation of Aeromonas species from meat and fish. Quantitative examination of samples may be performed with the same media, using a direct plating method. An incubation temperature of 25–30ºC is recommended for environmental and food samples.137 In general, the optimum recovery of aeromonads requires more than one agar medium. The same procedure may be used to isolate aeromonads from heavily contaminated clinical specimens such as feces, although in this case a temperature of incubation of 35ºC is recommended. BIBG medium was originally designed for the selective isolation of aeromonads from feces.141 Bile salts and brilliant green inhibit the growth of Gram-positive bacteria and Irgasan inhibits the growth of Gram-negative bacteria, which possess a type A nitratase. Bacteria that survive the selective process can be differentiated by their ability to assimilate-ferment xylose. Noninoculated BIBG is a purple-red, transparent medium. If xylose is fermented, acid is produced and the medium retains its original, noninoculated color. Mesophilic aeromonads are not able to ferment xylose and basic compounds are formed due to the breakdown of proteins, the medium changing from purple to a green-yellow color. Aeromonas species appear as translucent colonies and have a white-green appearance. The BIBG medium has been modified (mBIBG) by increasing the pH up to 8.7 and replacing xylose by soluble starch as a carbon source, with very good results.142 20.1.5.2  Biochemical Identification Once Aeromonas species have been isolated and detected, they must be identified at species level. The greatest difficulties in the identification of Aeromonas strains at species level derive from the current number of recognized taxa (n=14) and the lack of appropriate phenotypic schemes that allow each of these groups to be distinguished from the others.92 Most clinical microbiological laboratories use the Aerokey II system as an accurate and reliable system for the phenotypic identification of currently recognized Aeromonas species isolated from clinical specimens and related to human diarrhea, namely A. hydrophila, A. caviae, A. veronii biovar sobria, A. veronii biovar veronii, A. schubertii, A. jandaei, and A. trota.143 The Aerokey II system was used in a study of 167 clinical isolates Aeromonas strains and is based on the evaluation of 18 biochemical tests: Esculin hydrolysis, Voges-Proskauer reaction, pyrazinamidase activity, CAMPlike factor (aerobic only); fermentation of arabinose, mannitol, and sucrose; susceptibility to ampicillin, carbenicillin, cephalothin, and colistin; decarboxylation of lysine and ornithine; arbutin hydrolysis; and production of indole, H2S, gas from glucose, and hemolysis (TSA with 5% sheep erythrocytes). The Aerokey II system correctly identified 97% of the

60 coded clinical isolates from an independent laboratory and 100% of the reference strains at species level, and it may be considered as a very accurate Aeromonas identification system.143 However, Aerokey II and many other biochemical schemes144–146 used in clinical laboratories for aeromonad identification predate the description of newer taxa and may not be applicable for aeromonad identification described more recently. An interesting study has been performed on 193 strains of Aeromonas representing the 14 species currently recognized for 63 phenotypic traits.147 In this study, a schema was designed for the identification of aeromonads, first at group and then at species level within each group, and it recommended the tests to be performed in clinical and reference laboratories. After isolation of aeromonads on selective media, it is possible to identify clinical isolates of Aeromonas species in one of the three major complexes or groups of significance in human disease: the A. hydrophila group (A. hydrophila, A. bestiarum and A. salmonicida); the A. caviae group (A. caviae, A. media, and A. eucrenophila), and the A. sobria group (A. veronii HG8, A. jandei, A. schubertii, and A. trota). To separate species within complexes, sets of biochemical reactions were described.1,147

20.1.6  Molecular Detection and Identification Molecular methods such as nucleic acid amplification with the polymerase chain reaction (PCR) or genetic probes have been used profusely over the past 15 years for the direct detection of aeromonads in a variety of matrices such as raw milk,148 water,149,150 flies,151 fish,152 feces,153 and others.154 PCR methods are usually more sensitive than culturing for complex matrices that are heavily contaminated with other bacteria that interfere with culture detection. A. hydrophila has been detected in raw milk with a detection limit of 100 CFU/g, with a PCR detection rate of 23% and 14% for culturing.148 Further, selective media are supplemented with antibiotics that inhibit the growth of sensitive A. trota and some strains of A. caviae. A PCR method targeting the aerolysin gene has been described for the direct detection of Aeromonas species in feces. The PCR method detected the aerolysin gene in 68% of fecal samples as compared to 64% of aeromonads detected by conventional selective media. The Aeromonas species detected by PCR aerolysin gene amplification included A. hydrophila (55.8%), A. caviae (17.6%), A. veronii (10.2%), A. schubertii (4.4%), A. jandaei (2.9%) and A. trota (8.8%).155 PCR gene amplification has also been used for the detection of pathogenic bacteria and to differentiate pathogenic strains from non-pathogenic ones.156–158 A multiplex PCR was developed to detect hemolysin and aerolysin genes in A. hydrophila and A. sobria and used to screen many clinical and reference strains. Five genotypes were identified based on the presence or absence of ahh1, asa1, and aerA genes.159 The presence and number of hemolysin genes corresponded

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to cytotoxin titres when the phenotypic expression of toxin was evaluated in Vero cell culture cytotoxicity assays. PCR amplification of the A. hydrophila lip gene has been used successfully at our laboratory for the detection and identification of A. hydrophila HG1.160 We have reported that the PCR amplification of the aroA gene with specific primers can be used as a useful target gene for the detection and identification of all aeromonads at genus level.161 When PCR amplification of the aroA gene was combined with restriction fragment length polymorphism (RFLP) using the endonuclease Hae II, a different RFLP was obtained for each of the 14 reference Aeromonas strains used in this study, representing 14 Aeromonas species. Additionally, to facilitate the assay we did not exploit the possibility of using nucleotide sequence analysis of aeromonads aroA genes; however, this aroA housekeeping showed high genetic diversity and can be used in phylogenetic studies for Aeromonas species and strains identification. Microarray DNA probes have been used to study the population interactions of marine bacteria, where aeromonads represent a significant percentage of the microbiota.162 Other studies have applied microarray technology to identify A. hydrophila cytotoxic enterotoxin-inducing genes in macrophages, thus demonstrating the potential of microarrays for elucidating the intracellular mechanisms of pathogenesis.163,164 Microarrays and proteomics have also been used to study the effects of Aeromonas cytotoxic enterotoxin on human epithelial cells.165 Fluorescent in situ hybridization (FISH) is another probe technology that has been used to characterize aeromonads present in goldfish feces.166 In the last 15 years the molecular identification of aeromonads has been headed by increased use of 16S rRNA sequences.167–169 However, the use of this molecular marker induces a complication because ribosomal genes are organized in operons and bacteria may contain up to 15 copies of the ribosomal operon,170 and the evidence suggests that intragenomic heterogeneity exists, even in Aeromonas.55,171,172 A pioneer study to reconstruct the phylogeny of Aeromonas species revealed that A. trota and A. caviae only show three nucleotide differences in their 16S rRNA gene sequences but the DNA–DNA hybridization values for these species are 30%, indicating that they are distantly related. In contrast, A. veronii and A. sobria differ by 12 nucleotides but have a DNA–DNA hybridization value of 60–65%, indicating that they are closely related.173 This lack of congruence may be explained in terms of lateral gene transfer and recombination.174 This is why in the genus Aeromonas the 16S rRNA gene sequence is only useful to define isolates at genus level, it being necessary to use other single-copy and stable housekeeping genes for correct Aeromonas identification. Housekeeping genes are considered better molecular markers than the 16S rRNA genes for the study of taxonomic and phylogenetic relationships at species level, and the current consensus is that an informative level of phylogenetic data should be obtained from the determination of nucleotide sequences of a minimum of five housekeeping genes

Molecular Detection of Foodborne Pathogens

(genes under stabilizing selection for encoded metabolic functions).60 However, several phylogenetic studies addressing the genus Aeromonas are based on a lower number of housekeeping genes. Thus, the rpoD gene, encoding the σ70 factor of DNA-dependent RNA polymerase, was evaluated in comparison to gyrB (which encodes the β-subunit of DNA gyrase) for phylogenetic and taxonomic studies of 70 Aeromonas strains representing all known species of the genus with excellent results, especially for closely related taxa.175 Thirty-three Aeromonas strains isolated from human and animal specimens were characterized using 16S rRNA, gyrB, and rpoB (which encodes the β subunit of DNA-dependent RNA polymerase) gene sequences. In this study, it was concluded that 16S rRNA gene sequencing is useful to define isolates at the genus level; however, gyrB gene sequences are also a powerful tool for differentiating Aeromonas genospecies and even strains within species.56 rpoB gene sequences are more conserved than gyrB sequences, but the resolution of rpoB is sufficient to infer phylogenetic relationships and taxonomic identifications within Aeromonas.56 Recently, similar studies have been performed to update the phylogeny of the genus Aeromonas by including new described species, such as A. simiae, A. molluscorum, and A. culicicola.53 A novel phylogenetic marker—the dnaJ gene—complemented with DNA–DNA hybridization studies has been used to clarify some controversies within the genus Aeromonas.58 The dnaJ gene encodes a heat-shock protein 40 and has been reported to be a suitable target for phylogenetic study and the identification of species of Mycobacterium,176 Legionella,177 and Streptococcus.178 The mean dnaJ sequence similarity was 89.2%, indicating that divergence within the genus Aeromonas is comparable to that of gyrB and rpoD.175 At intraspecies level, the rate of nucleotide substitution did not exceed 3.3% except for three A. hydrophila subspecies, which showed far larger distances. At interspecies level, the rate of nucleotide substitution was greater than 5.2% for most Aeromonas strain pairs, but it was only 1.7–3.3% for a cluster including three species (A. veronii, A. allosaccharophila and A. culicicola) and another one that included A. echeleia and Aeromona sp. HG11. This study suggests that A. echeleia and Aeromona sp. HG11 belong to the same genetic group since they share 82–85% relatedness by DNA–DNA hybridization and only 1.3% divergence with the gyrB sequence.175 Based on genotypic and phenotypic evidence it has also been suggested that A. allosaccharophila and A. culicicola may be affiliated to A. veronii.175 Recently, the recA gene sequence has been evaluated for the classification of Aeromonas strains at genotype level, with similar results to those obtained when this gene was included in a multi locus sequence typing scheme (MLST).1,59,179 A. aquariorum sp. nov. has recently been described based on phylogenetic analysis, using 16S rRNA, gyrB, and rpoD gene sequences, complemented with DNA–DNA hybridization, MALDI-TOF MS analysis, and extensive biochemical and antibiotic susceptibility tests.180

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20.2  Methods 20.2.1  Reagents and Equipment Reagents Sterile distilled water 10 × PCR buffer MgCl2 dNTPs Primer sets Taq polymerase Agarose Gel electrophoresis buffer Ethidium bromide

Equipment DNA thermal cycler Gel electrophoresis apparatus Ultraviolet light box

20.2.2  Sample Collection and Preparation Clinical specimens, where Aeromonas species are usually present in high numbers, should be collected and transported according to recommended clinical laboratory practice.181 Collection and preparation of environmental and food samples may be processed according to the principles outlined by official agencies.182,183 Recovery of Aeromonas from clinical specimens such as stool samples should use a broth enrichment procedure. Stool specimens may be inoculated in APW and incubated overnight aerobically with shaking (100 rpm). One loopful of enriched sample may be plated onto xylose-deoxycholatecitrate agar (XDCA) and sheep-blood agar (5% sheep blood) supplemented with 30 µg/ml ampicillin (ASBA), followed by incubation at 37ºC overnight. Colorless colonies that do not ferment xylose on XDCA and ampicillin-resistant hemolytic colonies on ASBA should be tested for catalase and oxidase positivity and the resistance to the O129 vibriostatic agent. Presumptive Aeromonas colonies are ready for a phenotypic identification scheme.147 If a molecular identification method is going to be used, presumptive Aeromonas colonies are then suspended in 100 µl of TE buffer, vortexed for 1 min, and incubated at 96ºC for 10 min in a water bath. After centrifugation of the tube at 12,000 × g, the supernatant is transferred to a clean tube and 3–5 µl samples can be used as a DNA template for PCR assays. To recover Aeromonas from food samples, 25 g of each food sample are weighted and transferred aseptically to a sterile stomacher bag and homogenized using a Stomacher device for 1–2 min with 225 ml of APW. For Aeromonas quantitative (CFU/g) analysis 0.1 ml of ten-fold serial dilutions are plated, before enrichment, onto two selective culture media: Ampicillin-Dextin-Agar (ADA) and modified Bile Salts Irgasan-Brilliant Green (m-BIBG).142 Both media are incubated for 24 h at 30ºC and presumptive Aeromonas colonies enumerated. Aeromonas strains typically produce yellow colonies on ADA medium and purple colonies on mBIBG medium. Presumptive Aeromonas are ready for phenotypic or molecular identification schemes as commented above. Aeromonas qualitative (presence/absence per 25 g) analysis may require enrichment by incubation for 24 h at 30ºC prior

to inoculation onto ADA and mBIBG media. Presumptive Aeromonas colonies are identified as described above. Food samples are defined as being positive for Aeromonas if confirmed Aeromonas colonies are isolated on either ADA or mBIBG media.

20.2.3  Detection Procedures Principle. Molecular detection of Aeromonas may be more sensitive than culture for complex matrices with dense background microbiota. However, it is recommended to use an enrichment medium followed by selective media as described above. Presumptive Aeromonas colonies on selective media can be confirmed as Aeromonas species by using a very simple and easy molecular method consisting in PCR amplification of the aroA gene, which is being used in our lab with excellent results, although other molecular methods may provide good results as well,56,58,59 followed by restriction fragment length polymorphism (RFLP) analysis using HaeII endonuclease. The aroA gene is involved in a universal biosynthetic pathway for the biosynthesis of folic acid and aromatic amino acids, which is functional in all bacteria. The aroA gene nucleotide sequences is very well conserved within Gram-negative bacterial groups and may be used as a powerful and appropriate tool for the identification of bacteria to the genus level by the use of a simple and inexpensive procedure such as aroA PCR amplification. The aroA gene primers consist of a 24 nucleotides forward primer PF1 (5′-TTTGGAACCCATTTCTCGTGTGG C-3′) corresponding to positions 15–38 of the A. salmonicida aroA gene nucleotide sequence (from the GenBank data base under accession number L05002), and a 25 nucleotides reverse primer PR1 (5′-TCGAAGTAGTCCGGGAAGGTCT TGG-3′), which corresponds to positions 1253 to 1229. The combination of primers (PF1 and PR1) from A. salmonicida aroA gene enables the synthesis of an expected 1236-bp DNA fragment, which represents bulk of aroA gene sequence of all Aeromonas reference strains tested. Aeromonas spp. aroA PCR amplification may be performed from both extracted nucleic acids and bacterial cells; however, to prevent contaminations and avoid an excessive DNA manipulation, all data presented here were obtained from presumptive Aeromonas colonies, which simplified the procedure. Sensitivity was very high; amplification which resulted in detectable levels of PCR product was achieved when a minimum of eight CFU of any Aeromonas spp. were lysed, on the basis of an average of five repeated testing of viable cells and PCR assays. Procedure: (1) Prepare PCR mixture (50 µl) containing 1 µl of DNA-containing sample (extracted by the rapid boiling method above), 1.25 U of Taq DNA polymerase (Boehringer GmbH, Mannheim, Germany), 5 µl of 10× PCR amplification buffer (100 mM Tris-HCl, 20 mM MgCl2, 500 mM KCl [pH 8.3]), 1 µM each

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Figure 20.1  Agarose gel electrophoresis of PCR amplification products from Aeromonas spp. Lanes 1–13: A. eucrenophila CECT 4224, A. jandaei CECT 4228, A. salmonicida CECT 894, A. hydrophila CECT 839, A. sobria CECT 837, A. enteropelogenes CECT 4487, A. schubertii CECT 4241, A. ichthiosmia CECT 4486, A. media CECT 4232, A. allosaccharophila CECT 4199, A. caviae CECT 838, A. encheleia CECT 4341, and A. trota CECT 4255, respectively. M, standard DNA size marker φX174 HaeIII digested.

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Figure 20.2  Agarose gel electrophoresis of fragments produced by HaeII digestion of 13 species of Aeromonas aroA genes amplified by PCR. Lanes 1–13: reference Aeromonas strains ordered as in Figure 20.1. M, DNA molecular weight marker (100-bp ladder).

primer (PF1 and PR1), 0.5 mM ­deoxynucleosides triphosphate, and double-distilled water (to a final volume of 50 µl). To minimize evaporation, add 50 µl of mineral oil to the mixture. (2) Carry out the following cycling program on a DNA thermal cycler (Perkin-Elmer Cetus, Norwalk, CT or other machine): one cycle of 94ºC for 2 min; 40 cycles of 92ºC for 1 min, 50ºC for 1 min, and 72ºC for 1 min; and 1 cycle of 72ºC for 10 min. Keep the reaction mixture at 4ºC until analysis. (3) Run 15 µl of PCR products to confirm the expected 1236-bp DNA fragment on a 1% agarose gel (prepared in 0.5× TBE) containing 0.5 µg/ml ethidium bromide (Figure 20.1). (4) Digest 20 µl of PCR product with 5 U of HaeII restriction endonuclease for 2 h, and separate in 3% agarose gels (prepared in 0.5× TBE) containing 0.5 µg/ml ethidium bromide (Figure 20.2) Comments: The PCR primers, PF1, and PR1, from A. salmonicida aroA gene produce an expected 1236-bp DNA fragment, which represent most of aroA gene sequence of 13 Aeromonas reference strains (Figure 20.1). However, no PCR amplification product is obtained when members of Enterobacteriaceae, Vibrionaceae, and Pseudomonadaceae families’ cells are used as negative controls. A typical example of aroA gene

200 100

Figure 20.3  Agarose gel electrophoresis of fragments produced by HaeII digestion of Aeromonas aroA genes amplified by PCR. Lanes 1 and 2, environmental strains phenotypically identified as A. caviae; Lane 3, A. caviae CECT 838; Lanes 4 and 5, environmental strains phenotypically identified as A. media; Lane 6, A. media CECT 4232; Lane 7, A. sobria CECT 837; Lane 8, environmental strain phenotypically identified as A. sobria; Lanes 9–11, clinical isolates identified as A. salmonicida; Lane 12, A. salmonicida CECT 894.

PCR amplification of 13 Aeromonas strains is shown in Figure 20.1. The utility of this procedure is further enhanced when combined with RFLP of the amplified aroA gene, which results in specific patterns for each strain (Figure 20.2). This molecular method (aroA gene PCR-RFLP) is also used for identification of Aeromonas environmental isolates (two strains of A. caviae, two strains of A. media, and one strain of A. sobria) or clinical isolates (three strains of A. salmonicida from diseased fish) that are previously identified by biochemical characteristics.145 We found good correlation between reference strain RFLPs and Aeromonas isolate RFLPs (Figure 20.3) as confirmed by phenotypic identification as well.

20.3 Conclusions and Future Perspectives Members of the genus Aeromonas are Gram-negative, facultatively anaerobic, rod-shape bacteria (1–3.5 µm), although cocci may appear with diameters ranging from 0.3 to 1.0 µm. They are chemoorganotrophic, with an optimal growth temperature of 30ºC; a minimum growth temperature of 4ºC and a maximum growth temperature of 45ºC. They are oxidase- and catalase-positive, secrete many extracellular products (most of them considered virulence factors), reduce nitrate, ferment D-glucose, and are resistant to the vibriostatic agent O129. The natural habitat of Aeromonas is in aquatic environments, marine water and fresh water associated with aquatic animals and sediments. They are frequently isolated from foods and stools, and some are considered to be opportunistic pathogens of human beings and animals owing to the detection of suspected foodborne outbreaks.1,49 Aeromonas spp are motile through a polar flagellum, with the exception of A. salmonicida and A. media which are non motile species. Most Aeromonas spp are mesophilic, although A. salmonicida is a psychrophile.

283

Aeromonas

Most Aeromonas spp also produce an array of extracellular enzymes: proteases, DNAses, RNAses, lipases, lecithinases, amylases, enterotoxins, cytotoxins, and hemolysins.181 Aeromonas has been frequently misidentified at species level owing to the current number of recognized taxa, with continuous incorporations of new species and a lack of a panel of phenotypic properties that allow species to be discriminated or distinguished. The discrepancies of different sets of data using the same Aeromonas strains have contributed to the difficulties in the identification of these microorganisms as well as the use of select biochemical characteristics for newly described species or different conditions of manipulation at different laboratories.52,184–186 However, recent studies have presented cumulative biochemical data for many Aeromonas strains representing all taxa described that allow accurate identification of current Aeromonas species with the depth required at laboratory level.147 Recently, housekeeping gene sequence analysis (gyrB, rpoB, rpoD, danJ, or recA) complemented with DNA-DNA hybridization analysis and/ or 16S rRNA sequence analysis have provided new tools for the elucidation of Aeromonas taxonomy and have contributed to delineating closely related species.58,59,176–179 Although aeromonads have been known for 100 years, the first report of Aeromonas-due human infection was in 195413 and the first strain isolated from feces occurred in 1961.187 Since then, Aeromonas species have been associated with human diarrhea, although the evidence has been inconclusive because the criteria that could be used to solve this issue (Koch’s postulates, an animal model reproducing human disease, and the failure of studies with human volunteers) have been insufficient and hence the debate remains open. Three Aeromonas species (A. hydrophila HG1, A. caviae HG4, and A. veronii biovar sobria HG8) have been isolated from human feces with significant frequency, representing 85% of isolates.188 A. trota HG14, A. jandei HG9, and A. schubertii HG12 are rare fecal isolates73. If Aeromonas enteric infections are transmitted by food, drinking water or person-to-person contact, and symptomatic infections are strain-specific, it would be expected that the same strain would be isolated in patients and in foods, in drinking water or in an Aeromonas-transmitting person. However, this is not the general situation,68 and only two cases have been reported that suggest transmission: one from food to a patient with Aeromonas-associated diarrhea,70 and another one from person to person.71 In both studies, ribotyping was used to demonstrate identity. Because of the abundance of Aeromonas species in nature and their potential involvement in human and animal disease, further investigation is needed to detect virulence mechanisms in sets of Aeromonas strains associated with human and animal disease. This solution would be of considerable importance for clinicians and for clinical microbiology laboratories with a view to investigating the presence of Aeromonas in clinical specimens. It would also be very important for public authorities to regulate specific microorganisms that may be found in drinking water, as well as food production, distribution, and consumption guidelines.

References













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Aeromonas 149. Kong, R.Y. et al. Rapid detection of six types of bacterial pathogens in marine waters by multiplex PCR. Water Res., 36, 2802, 2002. 150. Peng, X.X. et al. Immuno-capture PCR for detection of Aeromonas hydrophila. J. Microbiol. Methods, 49, 335, 2002. 151. Nayduch, D. et al. Detection of Aeromonas caviae in the common housefly Musca domestica by culture and polymerase chain reaction. Epidemiol. Infect., 127, 561, 2001. 152. Gonzalez-Rodriguez, M.N. et al. PCR detection of potentially pathogenic aeromonads in raw and cold-smoked freshwater fish. J. Appl. Microbiol., 93, 675, 2002. 153. Fukushima, H., Tsunomori, Y. and Seki, R. Duplex real-time SYBR green PCR assays for detection of 17 species of foodor waterborne pathogens in stools. J. Clin. Microbiol., 41, 5134, 2003. 154. Ji, N. et al. Universal primer PCR with DGGE for rapid detection of bacterial pathogens. J. Microbiol. Methods, 57, 409, 2004. 155. Kannan, S. et al. Direct detection of diarrheagenic Aeromonas from faeces by polymerase chain reaction (PCR) targeting aerolysin toxin gene. Eur. Rev. Med. Pharm. Sci., 5, 91, 2001. 156. Kingombe, C.I. et al. The usefulness of molecular techniques to assess the presence of Aeromonas spp. harboring virulence markers in foods. Int. J. Food Microbiol., 94, 113, 2004. 157. Kingombe, C.I. et al. PCR detection, characterization, and distribution of virulence genes in Aeromonas spp. Appl. Environ. Microbiol., 65, 5293, 1999. 158. Sen, K. and Rodgers, M. Distribution of six virulence factors in Aeromonas species isolated from U.S. drinking water utilities: a PCR identification. J. Appl. Microbiol., 97, 1077, 2004. 159. Wang, G. et al. Detection and characterization of the hemolysin genes in Aeromonas hydrophila and Aeromonas sobria by multiplex PCR. J. Clin. Microbiol., 41, 1048, 2003. 160. Cascon, A. et al. Identification of Aeromonas hydrophila hybridization group 1 by PCR assays. Appl. Environ. Microbiol., 62, 1167, 1996. 161. Cascon, A. et al. RFLP-PCR análisis of the aroA gene as a taxonomic tool for the genus Aeromonas. FEMS Microbiol. Lett., 156, 199, 1997. 162. Stine, O.C. et al. Characterization of microbial communities from coastal waters using microarrays. Environ. Monit. Assess., 81, 327, 2003. 163. Galindo, C.L. et al. Microarray analysis of Aeromonas hydrophila cytotoxic enterotoxin-treated murine primary macrophages. Infect. Immun., 72, 5439, 2004. 164. Galindo, C.L. et al. Identification of Aeromonas hydrophila cytotoxic enterotoxin induced genes in macrophages using microarrays. J. Biol. Chem., 278, 40198, 2003. 165. Galindo, C.L. et al. Microarray and proteomics analyses of human intestinal epithelial cells treated with the Aeromonas hydrophila cytotoxic enterotoxin. Infect. Immun., 73, 2628, 2005. 166. Asfie, M., Yoshijima, T. and Sugita, H. Characterization of the goldfish fecal microflora by the fluorescent in situ hybridization method. Fisheries Sci., 69, 21, 2003. 167. Borrell, N. et al. Identification of Aeromonas clinical isolates by restriction fragment length polymorphism of PCR-amplified 16S rRNA genes. J. Clin. Microbiol., 35, 1671, 1997. 168. Figueras, M.J. et al. Extended method for discrimination of Aeromonas spp. by 16S rDNA RFLP analysis. Int. J. Syst. Evol. Microbiol., 50, 2069, 2000. 169. Graf, J. Diverse restriction fragment length polymorphism patterns of the PCR-amplified 16S rRNA genes in Aeromonas veronii strains and possible misidentification of Aeromonas species. J. Clin. Microbiol., 37, 3194, 1999.

287 170. Klappenbach, J.A., Dunbar, J.M. and Schmidt, T.M. rRNA operon copy number reflects ecological strategies of bacteria. Appl. Environ. Microbiol., 66, 1328, 2000. 171. Boucher, Y. et al. Intragenomic heterogeneity and intergenomic recombination among haloarchaeal rRNA genes. J. Bacteriol., 186,3980, 2004. 172. Coenye, T. and Vandamme, P. Intragenomic heterogeneity between multiple 16S ribosomal RNA operons in sequenced bacterial genomes. FEMS Microbiol. Lett., 228, 45, 2003. 173. Martinez-Murcia, A.J., Benlloch, S. and Collins, M.D. Phylogenetic interrelationships of members of the genera Aeromonas and Plesiomonas as determined by 16S ribosomal DNA sequencing: lack of congruence with results of DNADNA hybridizations. Int. J. Syst. Bacteriol., 42, 412, 1992. 174. Sneath, P.H.A. Evidence from Aeromonas for genetic crossing-over in ribosomal sequences. Int. J. Syst. Bacteriol., 43, 626, 1993. 175. Soler, L. et al. Phylogenetic analysis of the genus Aeromonas based on two housekeeping genes. Int. J. Syst. Evol. Microbiol., 54, 1511, 2004. 176. Takewaki, S. et al. Nucleotide sequence comparison of the mycobacterial dnaJ gene and PCR restriction fragment length polymorphism analysis for identification of mycobacterial species. Int. J. Syst. Bacteriol., 44, 159, 1994. 177. Liu, H. et al. Use of the dnaJ gene for the detection and identification of all Legionella pneumophila serogroups and description of the primers used to detect 16S rDNA gene sequences of major members of the genus Legionella. Microbiol. Immunol., 47, 859, 2003. 178. Itoh, Y. et al. dnaJ and gyrB gene sequence relationship among species and strains of genus Streptococcus. Syst. Appl. Microbiol., 29, 368, 2006. 179. Carnahan, A.M. Genetic relatedness of Aeromonas species based on the DNA sequence of four distinct genomic loci. PhD Dissertation, University of Maryland, College Park, MD, 2001. 180. Martínez-Murcia, A.J. et al. Aeromonas aquariorum sp. nov., isolated from aquaria of ornamental fish. Int. J. Syst. Evol. Microbiol., 58, 1169, 2008. 181. Altwegg, M. Subtyping methods for Aeromonas species. In: Austin, B. et al. (eds), The Genus Aeromonas. John Wiley & Sons, New York, NY, 1996. 182. APHA. Standard Methods for Examination of Water and Wastewater, 20th edn. Washington, DC, 1998. 183. USFDA. Bacterial Analytical Manual Online http://www. cfsan.fda.gov/~ebam/bam-mm.html, 2001. 184. Esteve, C., Gutierrez, M.C. and Ventosa, A. DNA relatedness among Aeromonas allosaccharophila strains and DNA hybridization groups of the genus Aeromonas. Int. J. Syst. Bacteriol., 45,390, 1995. 185. Huys, G., Kämpfer, P. and Swings, J. New DNA-DNA hybridization and phenotypic data on the species Aeromonas ichthiosmia and Aeromonas allosaccharophila: A. ichthiosmia Shubert et al. 1990 is a later synonym of A. veronii HickmanBrenner et al. 1987. Syst. Appl. Microbiol., 24, 177, 2001. 186. Martínez-Murcia, A.J. et al. Phenotypic, genotypic and phylogenetic discrepancies to differentiate Aeromonas salmonicida and Aeromonas bestiarum from each other. Int. Microbiol., 8, 259, 2005. 187. Lautrop, H. Aeromonas hydrophila isolated from human feces and its possible pathological significance. APMIS, 51/S 144,299, 1961. 188. Moyer, N.P. et al. Media and methods for isolation of aeromonads from fecal specimens, A multilaboratory study. Experientia, 47, 409, 1991.

21 Arcobacter Kurt Houf

Ghent University

Contents 21.1 Introduction.................................................................................................................................................................... 289 21.1.1 The Genus Arcobacter...................................................................................................................................... 290 21.1.2 Arcobacter in Humans...................................................................................................................................... 290 21.1.3 Virulence Factors.............................................................................................................................................. 291 21.1.4 Antimicrobial Susceptibility............................................................................................................................. 292 21.1.5 Arcobacter in Animals..................................................................................................................................... 292 21.1.5.1 Arcobacter in Cattle......................................................................................................................... 292 21.1.5.2 Pigs as an Important Arcobacter Reservoir..................................................................................... 293 21.1.5.3 Arcobacter in Poultry...................................................................................................................... 293 21.1.5.4 Arcobacter in Other Animals.......................................................................................................... 294 21.1.6 Arcobacter in Food of Animal Origin.............................................................................................................. 294 21.1.7 Identification and Characterization of Arcobacter........................................................................................... 296 21.1.7.1 Molecular Identification................................................................................................................... 296 21.1.7.2 Molecular-based Characterization................................................................................................... 298 21.2 Methods.......................................................................................................................................................................... 298 21.2.1 Arcobacter Isolation.......................................................................................................................................... 298 21.2.2 Sample Preparation........................................................................................................................................... 298 21.2.3 Detection Procedures........................................................................................................................................ 299 21.2.3.1 Genus-specific Identification by PCR.............................................................................................. 299 21.2.3.2 Species-specific Identification by PCR............................................................................................ 299 21.2.3.3 Species Identification by PCR-RFLP.............................................................................................. 300 21.2.3.4 Characterization by ERIC-PCR....................................................................................................... 300 21.3 Conclusions and Future Perspectives............................................................................................................................. 300 References.................................................................................................................................................................................. 301

21.1  Introduction Bacteria, now known as campylobacters, were first isolated at the beginning of the twentieth century. In 1913, McFadyean and Stockman isolated Vibrio-like organisms from aborted ovine fetuses.1 Five years later, Smith described the isolation of Vibrio/Spirillum-like organisms from aborted bovine fetuses, which he considered the same species as described by McFadyean and Stockman.2 Although the cell shape of the those organisms was characteristic of the ­family Vibrionaceae, they failed to ferment carbohydrates, and were therefore transferred into a new genus, Campylobacter.3 Seventy years later, Ellis et al.4,5 and Higgins and Degre6 reported the isolation of aerotolerant, spiral-shaped Campylobacter-like organisms from the organs of porcine and bovine fetuses. Examination of all Campylobacter and Campylobacter-like isolates revealed two distinct biochemical groups: the group 1 isolates were identified as C. fetus, whereas the group 2 isolates were considered as the “aerotolerant campylobacters.”7,8 In a comprehensive study in 1985, Neill et al.9 provided a thoroughly documented description for a new species,

C. cryaerophila to include these ­“aerotolerant ­campylobacters” but emphasized also their phenotypic heterogeneity. In 1988, Thompson et al.10 showed by partial 16S rRNA sequence analysis that C. cryaerophila and C. nitrofigilis, an organism isolated from the roots of Spartina alterniflora, exhibited more homology with each other than with other campylobacters, suggesting that classification of both C. cryaerophila and C. nitrofigilis in another genus would be appropriate. In the early 1990s, Kiehlbauch et al.11 found two subgroups among C. cryaerophila strains by phenotypic characterization and DNA-DNA hybridization. Catalase positive strains that were able to grow under aerobic conditions at 30°C but not in a medium with glycine or on MacConkey agar, were considered as C. cryaerophila. The name C. butzleri was proposed for the aerotolerant isolates that were negative to weakly catalase positive, and for which growth was observed in glycine minimal medium an on MacConkey agar. In 1991, after a polyphasic taxonomic study, Vandamme et al.12 transferred C. cryaerophila and C. nitrofigilis into a new genus, Arcobacter. Subsequently, C. butzleri was reclassified as 289

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A. butzleri and A. skirrowii was proposed for yet another group of animal associated Arcobacter strains.13

21.1.1  The Genus Arcobacter Due to the close phenotypic and genotypic affiliation, the genus Arcobacter was classified together with the genus Campylobacter into the family Campylobacteraceae.14 Together with the genera Helicobacter, Wolinella, and Sulfurospirillum, they constitute the most important representatives of a distinct group referred to as the rRNA superfamily VI, or as the ε-division of the class Proteobacteria.12 Arcobacter are Gram-negative, nonspore-forming, motile, spiral-shaped rods (0.2–0.8 × 0.5–5 µm), that are able to grow microaerobically or aerobically.12 They have the ability to grow from 15 to 37°C, with an optimal growth in microaerobic conditions at 30°C. The growth at 15°C and the aerotolerance are the features that differentiate Arcobacter from Campylobacter species.12,15 In 2007, the complete genome sequence analysis of A. butzleri revealed however that the majority of its proteome is most similar to those of Sulfuromonas denitrificans and Wolinella succinogenes, and to those of the deep-sea vent Epsilonproteobacteria Sulfurovum and Nitratiruptor.16 The presence of pathways and loci associated with nonhost-associated organisms, as well as genes associated with virulence, suggests that A. butzleri is a free-living, waterborne organism.16 To date, six species have been described: A. butzleri, A. cryaerophilus, A. skirrowii, A. cibarius, A. nitrofigilis and A. halophilus.12,17–19 Predominantly A. butzleri has been associated with human infection as it has been isolated from stool of patients with diarrhea,20,21 but not from healthy humans.22 Furthermore, the presence of virulence genes,16 and its cytopathogenic effect on in vitro cell lines,23,24 resulted in the ­classification of this species as an emerging pathogen.25 Besides humans, A. butzleri has been isolated from healthy and ill live stock,26 poultry,27,28 nonhuman primates,29–31 and diverse environmental matrices such as water and sludge.32 A. cryaerophilus is a genotypically heterogeneous species, which was originally divided into two subgroups by fatty acid analysis13 or by restriction fragment length polymorphism (RFLP)-DNA analysis.33 As for A. butzleri, A. cryaerophilus strains have been isolated from a whole range of different matrices, including from stool of healthy humans22 as well as from patients with enteritis.20 A. skirrowii has first been isolated from feces of lambs with diarrhea,13 but later on also from preputial fluids of bulls, organs of bovine, porcine, and ovine aborted fetuses, animal feces, food, and water. Association of A. skirrowii with human disease has only rarely been reported.34,35 In 2005, A. cibarius was described as a new Arcobacter species, with A. cryaerophilus as the closest phylogenetic neighbor.18 The first representatives were isolated from broiler carcasses,18 but association with pigs has been suggested as well.36 Another potential new species has been isolated from the organs of aborted piglets and from duck feces,37,38 for which the name A. t­hereius sp. nov. will be proposed (personal communication).

Molecular Detection of Foodborne Pathogens

Besides the animal and human related species, different free-living, environmental Arcobacter species have been described. A. nitrofigilis is a nitrogen-fixing bacterium isolated from the roots of the salt marsh plant Spartina alterniflora.17 The strains prefer microaerobic growth conditions, but aerobic growth is possible in adapted culture media.39 An obligate microaerophile marine sulfide-oxidizing autotrophic bacterium has been described which produced hydrophilic filamentous sulfur as a novel metabolic end product for which the name Candidatus Arcobacter sulfidicus sp. nov. has been proposed.40 A phylogenetic affiliation of a Gramnegative bacterium isolated from water collected at a hypersaline lagoon on Hawaiian Islands to the genus Arcobacter was confirmed for which the name A. halophilus was introduced.19 Cells are slightly curved, obligate halophile and growth is observed in culture media containing 2–4% salt or 0.1% potassium nitrate under aerobic and microaerobic conditions at room temperature and at 37°C, and under anaerobic conditions at 37°C. Finally, DNA from a number of Arcobacter-like organisms has been detected in diverse sources as salt-water lakes, coastal seawater, oil wells, biocatalytic calcification reactor, sediments in the Black Sea, sludge, and production waters.41–52

21.1.2  Arcobacter in Humans Interest for Arcobacter in veterinary and human public health enhanced since the first report of the isolation of Arcobacter from food of animal origin.53 Since then, studies worldwide have reported the occurrence of Arcobacter on food and have highlighted the possible transmission to the human population. However, since the clarification of the taxonomic position of Arcobacter, only a few surveys have dealt with the clinical course of Arcobacter infection in humans. In 2000, Engberg et al.54 isolated one A. butzleri and one A. cryaerophilus from a total of 3267 clinical stool specimens. In the same year, Lastovica et al.55 reported an A. butzleri prevalence of 0.39% in a study realized on 19,535 diarrheal stool of pediatric patients. During an 8-year study period, Vandenberg et al.20 reported A. butzleri as the fourth most common Campylobacter-like organism isolated from 67,599 stool specimens form patients with diarrhea. In the study, A. butzleri was more associated with a persistent and watery, and less with bloody diarrhea compared with C. jejuni infection and the organism has been recovered from patients of different ages (<1–90 years old). Similar results have been reported from a surveillance network in France in 2006.21 Most clinical Arcobacter infections are single cases with the source of infection rarely identified. The first association of Arcobacter with human infection was reported in 1987. From the feces of a 35-year-old man with acute diarrhea and abdominal pain, A. cryaerophilus was isolated without detection of other pathogens.56 Since then, A. cryaerophilus, and A. butzleri infections were reported several times from stool samples of patients with acute diarrhea.11,35,54,57–60 In 2004, the first human isolation of A. skirrowii was reported by Wybo et al.34 from a 73-year-old patient with chronic

Arcobacter

diarrhea. More recently, A. skirrowii has been detected in diarrheal stool samples in South Africa.35 Besides the reports of the single cases of Arcobacterassociated enteritis, two Arcobacter outbreaks have been described. The first outbreak occurred in an Italian nursery and primary school where children suffered from recurrent abdominal cramps without diarrhea. The isolates were identified as A. butzleri and the identical phenotypic characteristics and genotypic profiles of the isolates suggested an epidemiological relation for all cases.60 Person-to-person transmission was assumed as the ongoing cause of infection. A second outbreak appeared during a scout camp, where 94 girls suffered from nausea, vomiting, abdominal cramps, and diarrhea.61 The outbreak was assumed to be correlated with the breakdown of the automated water chlorinating system. No clear cause of infection could be identified by the examination of stool samples, but A. butzleri was the only pathogen detected in the drinking water. Besides the correlation with gastro-enteritis, Arcobacter has also been implicated in extra-intestinal invasive diseases. Arcobacter butzleri was isolated from the blood of a neonate and the clinical data indicated an in utero sepsis.62 Yan et al.63 reported an A. butzleri isolation from two blood cultures of a man with liver cirrhosis displaying high fever and esophageal variceal bleeding, and in another bacteremia report, mentioned beside Escherichia coli and Streptococcus milleri, also the isolation of A. butzleri from a patient with an acute gangrenous appendicitis.59 Bacteremia due to A. cryaerophilus has been reported in a patient with hematogenous pneumonia,64 and in a traffic accident victim.65 The significance of Arcobacter as a cause of human diarrhea is still largely unknown, since clinical samples are not routinely tested for Arcobacter species. The symptoms are similar to those of Campylobacter infections, and Arcobacter infections often have a spontaneous recovery.66 This makes prevalence-determination difficult and mostly incorrect.20,67

21.1.3  Virulence Factors To assess the pathogenicity of Arcobacter for humans and animals, evaluation of potential virulence factors is required. However, up to now, little is known about the mechanisms of pathogenicity. A necessary state in the successful colonization, establishment, and ultimately production of disease by microbial pathogens is the ability to adhere to host surfaces such as mucous membranes, gastric, and intestinal epithelial or endothelial tissue. Therefore it is a common trait of microbial pathogens to express adherence factors responsible for recognizing and binding to specific receptor moieties of cells, thus enabling the bacteria to resist host strategies that would impede colonization. Since campylobacterioses and A. butzleri-related illnesses have similar clinical features, it might be expected that some C. jejuni virulence factors would be present in Arcobacter. In the genomic sequence analysis of A. butzleri strain LMG 10828T, homologs of the fibronectin binding proteins CadF

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and Cj1349, the invasin protein CiaB, the virulence factor MviN, the phospholipase PldA and the TlyA hemolysin were detected.16 Several other Campylobacter virulence-­associated genes however were not present with most notably the genes encoding the cytolethal distending toxin (CDT), which correlated with the study by Johnson and Murano who were unable to detect CDT-genes in Arcobacter species by PCR.68 Furthermore, three additional putative virulence determinants have been identified: irgA, which encodes an ironregulated outer membrane protein, hecA, a member of the filamentous hemagglutinin family, and hecB, a related hemolysin activation protein.16 The role and the functionality of these virulence determinants has not been determined yet. The first in vivo study with Arcobacter strains intravenous or intraperitoneal inoculated in rodents was unsuccessful.6 No clinical symptoms were present and lesions were not observed by autopsy. However, the invasion and colonization capacity has later been demonstrated by Wesley et al.69 as A. butzleri was isolated from feces and different organs up to seven days after experimental inoculation of piglets. Also the invasive capacity of A. cryaerophilus has been studied in an experimentally infected rainbow trout with death and clinical abnormalities reported.70 Agglutination of A. butzleri strains with human, rabbit, and sheep erythrocytes revealed the presence of adhesion molecules in Arcobacter.71 No fimbriae or pili were observed by electron microscopic examination, but an immunogenic hemagglutinin of 20 kDa was characterized. The hemagglutinin consisted of a lectin-like molecule, sensitive to heat treatment and enzymatic proteolysis, which could interact with a D-galactose-containing erythrocyte receptor. Several in vitro studies on the adhesion, invasion, ­interleukine-8- and toxin-production capacity by Arcobacter strains of different origin on Vero-, Hep-2, INT407, Caco-2, IPI-2I, and HeLa-cells have been performed.22–24,72,73 A strong cytopathogenic effect was observed in Vero-cells comparable to the effect of VT-toxin of E. coli O157,72,74 whereas Hep-2- and HeLa-cells showed weak cytopathogenic liason. Cell rounding and nuclear pyknosis was observed with Arcobacter isolates on HeLa and Intestine 407 cells and cell elongation on CHO-cells.24 The presence of a vacuole-forming toxin, different from Campylobacter CDT, in Arcobacter strains has been demonstrated in a Vero cell culture set-up.75 Fernandez et al.73 and Carter76 performed in situ studies about the existence of toxigenic and invasive capacities of Arcobacter. In those studies, the toxigenic capacity of Arcobacter from animal origin was determined in the rat and rabbit and pig ileal loop tests, respectively. In both studies, distention of the ileal loops with fluid accumulation and enhanced electrolyte concentrations was observed. The information based on molecular and in vitro studies, presently available suggest that adhesion, invasion and toxin production could be mechanisms by which Arcobacter species may cause disease. However, whereas colonization of the intestine and production of cytotoxins by bacteria is generally associated with bloody diarrhea, this symptom is rarely described in the human cases reported to date. Further

292

studies are certainly necessary to elucidate the pathobiology of Arcobacter species.

21.1.4  Antimicrobial Susceptibility Specific standardized procedures for susceptibility testing of Campylobacteraceae and resistance breakpoints have not been established. Consequently, a number of different testing methods, such as agar and broth (micro)dilution,77–79 disc diffusion,80–82 and the E-test 63,83 have been used for Arcobacter susceptibility testing in clinical, veterinary, and food microbiology. Furthermore, due to the fastidious nature and the microaerobic growth requirements of those microorganisms, the quality control limits given for nonfastidious organisms in aerobic atmosphere are not adequate.84 It is known that an increased level of carbon dioxide does decrease the effect of certain antimicrobials such as macrolides and fluoroquinolones. This will certainly occur in the microaerobic atmosphere required for the growth of Campylobacteraceae, and should be taken into account when interpreting susceptibility patterns.84 Comparison of the broth microdilution, the E-test and the agar dilution method showed overall comparable results for Campylobacter susceptibility testing when performed under the same microaerobic conditions and incubation temperature. Especially the minimum inhibitory concentration (MIC) obtained by the three methods of ciprofloxacin and erythromycin were in accordance with each other.85 In 1992, Kiehlbauch et al.79 applied the broth microdilution technique under aerobic atmosphere for susceptibility testing of the same panel of antimicrobials for A. butzleri and A. cryaerophilus isolates. The MICs for ciprofloxacin, erythromycin, doxycycline, and nalidixic acid differ by no more than one dilution from those obtained in the study of Houf et al.86 However, with gentamicin, MICs for A. butzleri and A. cryaerophilus ranged from ≤0.12 to 0.5 µg/ml, whereas in the study of Houf et al.86 the MICs ranged from 0.5 to 4 and 0.25 to 2 µg/ml, respectively. It is unknown whether the higher carbon dioxide concentration may decrease the activity of aminoglycosides as well. In the susceptibility study by Fera et al.77 even higher MIC ranges for both ciprofloxacin and gentamicin were obtained, although the same incubation atmosphere and temperature were applied. Whether the different origin of the strains in the studies, water versus human stools and poultry carcasses, is the cause of those MIC shifts, is not clear. In contrast to thermophilic Campylobacter species, of which some strains demonstrate resistance to quinolones and cross-resistance between nalidixic acid and quinolones, most of the Arcobacter strains seem to be susceptible to both antimicrobial agents. Remarkable is however the lowered susceptibility and even resistance to ciprofloxacin of A. butzleri strains isolated from poultry.86 The latter finding is also demonstrated by the concentrations required to inhibit growth of 50% of the strains (MIC50): the MIC50 for A. butzleri isolated from poultry is 0.12 whereas the MIC50 for human strains is 0.03.86 The use of fluoroquinolones for treatment of poultry may be the basis for this decreased susceptibility.

Molecular Detection of Foodborne Pathogens

Since particular poultry products are incriminated in the transmission of Arcobacter to humans, the presence of ­antimicrobial-resistant Arcobacter species in fresh poultry products can have public health implications. Today, data indicate that some Arcobacter isolates from poultry products are resistant and that multidrug resistance occurs. Especially the resistance to erythromycin and the decreased susceptibility to ciprofloxacin may have human health implications, as the two antimicrobials are generally prescribed as first-line drugs for the treatment of infections with Campylobacteraceae.

21.1.5  Arcobacter in Animals Apart from A. nitrofigilis and A. halophilus, Arcobacter species have been incriminated with various animal diseases including abortion, septicemia, mastitis, gastritis, and enteritis.11,29,87–91 However, several studies reported the occurrence of Arcobacter in healthy livestock and poultry, detected by different isolation methods and molecular techniques.26,92–99 Beside the clinical relevance, the occurrence of Arcobacter in healthy animals may act as significant reservoir and infection source to humans. 21.1.5.1  Arcobacter in Cattle In cattle, Arcobacter have been associated with pathologies such as mastitis and reproduction disorders, but have more commonly been isolated from feces of clinically healthy animals.93,95 In 1977, Arcobacter were isolated for the first time from the placenta and the internal organs of aborted bovine fetuses.4 Although several studies have confirmed these observations,6,100,101 no studies have clearly identified Arcobacter as a causative agent of disease in cattle. Moreover, Arcobacter have been isolated from preputial sheath washing samples of bulls with no association of breeding problems in the herd,102 and detected in vaginal swabs without reproduction problems in the cows.98 Besides the association with reproduction abnormalities, there are also reports about Arcobacter-associated mastitis. Arcobacter has been isolated from raw milk samples of freshly calved dairy cows during a mastitis outbreak, characterized by the presence of fine granular clots and very high cell counts in the milk.90 The isolate was used for an experimental inoculation of the udder of young dairy cows, which developed an acute clinical mastitis in the inoculated quarters. The challenge organism was only recovered from one of the cows, in a milk sample taken 4 h after inoculation. Beside the association with disease, Arcobacter have been detected in the feces of healthy cows, and the occurrence in the gut of healthy cows involves a potential risk of contamination of the environment and the human food chain, with carcass contamination caused by fecal contamination during the slaughter process. The occurrence of Arcobacter in cattle at different stage of production has been studied by several researchers and is listed in Table 21.1. The prevalence reported range from 2% to 39.2%, depending on the trail design (number of samples, sampling technique), country, season, age, and type of animals (calf or adult; dairy or fattening), isolation methods, and on-farm risk factors.

293

Arcobacter

Table 21.1 Prevalence of Arcobacter Species in Cattle Country

Sampling Place

USA USA

Farm Farm

Japan Turkey Belgium

Slaughterhouse Slaughterhouse Farm Slaughterhouse

Animal Type Dairy cows Fattening cattle Dairy cows Calves Cattle Fattening cattle Cattle Fattening cattle

In the studies, A. butzleri, A. cryaerophilus and A. skirrowii have been recovered with A. butzleri as the most frequently isolated species,95,97–99 except in the study of Van Driessche et al.,93 where A. cryaerophilus was the dominant species. Differences in physiological condition, housing, and feeding between calves, dairy cows and fattening cattle has an influence on the presence of Arcobacter, since different prevalence were observed between different animal groups.93,97 Of the calves, only 2% of the animals excreted Arcobacter whereas for fattening and dairy cattle, a prevalence up to 39% has been reported. 21.1.5.2  Pigs as an Important Arcobacter Reservoir As in cattle, the first reports of Arcobacter in pigs were associated with reproduction disorders. In several studies, Arcobacter have been isolated from the placenta and the internal organs of aborted fetuses with more late terms abortions, repeat breeding and a higher than usual rate of stillbirths observed on the farms.5,6,88 Antibiotic treatment or auto-vaccinations had none or only little improvement. At present, no routine screening for Arcobacter in pig farms has been established as the occurrence of abortions, whether caused by Arcobacter or not, seldom exceeds the reproduction parameters. However, results of a Danish survey suggest that the role of Arcobacter as etiological agent of abortion in pigs should not be ignored.88 In response to the Arcobacter associated abortus, some studies have dealt with the occurrence of Arcobacter in sows and boars. Arcobacter have been detected in oviductal tissues and uteri samples of sows with reproduction problems, and of sows with vaginal discharge.5,88,90 On farms with a history of Arcobacter associated abortions, sows with reproduction problems expressed high antibody serum titers in a microscopic agglutination test.103 However, Arcobacter have also been isolated from vaginal swabs of sows without reproduction disorders.98 On farms with and without reproduction problems, Arcobacter have been recovered from preputial swabs of boars, but not from the semen,103 though experimentally Arcobacter infected semen induced a decrease of the conception rates in sows.104 In one study, Arcobacter have been isolated from pig stomach samples, but their role in the etiology of gastric ulcers is not clear.91 Arcobacter do not always cause pathologies in pigs, since they have been isolated many times from the feces

Number of Animals

Prevalence

1682 50 50 100 332 200 276 51

14.3 14.0 18.0 2.0 3.6 9.5 11.0 39.2

Reference 95 97

98 99 26,93

of clinically healthy animals (Table 21.2). In Japan, A. butzleri was the most frequently isolated species in the feces at slaughterhouse level, followed by A. cryaerophilus.98 In a epidemiological study in which the occurrence of Arcobacter in animals of different age was followed, both A. butzleri and A. cryaerophilus were isolated, and characterization by pulsed field gel electrophoresis revealed the presence of different strains, suggesting colonization of animals by multiple parent genotypes that may undergo genomic rearrangements during successive passages through the animals, or colonization with genotypes originating from different sources.96 In a Dutch study, in which the transmission routes in sows and their offspring on a breeding farm was followed, ­intra-­uterine transmission was demonstrated, with A. skirrowii as the most prominent species. Furthermore, follow-up of the piglets suggested a postnatal infection from the sows, newcomers or from the environment, resulting in a demonstration of both vertical and horizontal transmission.105 The pathogenicity of Arcobacter in pigs has been studied in vivo only once. After intraperitoneal inoculation of neonatal piglets, no lesions or clinical symptoms were observed and no Arcobacter were isolated from the organs post mortem. In a second trial, the effect of an infection per os in caesarean-derived and colostrum-deprived piglets was determined. A. butzleri were isolated from the feces for up to 10 days as well as from the lung, liver, kidney, ileum, and the brain.69 The intestinal colonization and multiplication of A. butzleri was demonstrated, in contrast with A. cryaerophilus and A. skirrowii, for which only a short duration of fecal shedding was recorded with no isolations from the organs. This may suggest the failure of those species to penetrate the intestinal barrier. 21.1.5.3  Arcobacter in Poultry No association of Arcobacter with pathologies in poultry has been reported. Nevertheless, there are conflicting reports in literature whether or not Arcobacter is part of the poultry intestinal flora.94,95,107–111 In most of the studies, Arcobacter have not been isolated from cecal content nor from litter or the feathers,27,83,95,110 though some studies reported the isolation of Arcobacter from cloacal swab samples.37,99 In contrast with turkeys, infection experiments of chicks with A. butzleri were not successful.108,112,113 Since only some Arcobacter

294

Molecular Detection of Foodborne Pathogens

Table 21.2 Prevalence of Arcobacter Species in Pigs Country USA USA

Sampling Place Slaughterhouse Farm 1 Farm 2 Farm 3

Japan Belgium The Netherlands

Slaughterhouse Slaughterhouse Farm Farm

Animal Type Market-age pigs Piglets Weaned pigs Sows Piglets Sows Piglets Pigs Sows Market-age pigs Market-age pigs Pigs Sows Piglets at birth Piglets of 2 weeks Piglets of 3 weeks

strains grow at 41°C, it is feasible that the high body temperature of birds inhibited or suppressed Arcobacter growth and colonization. The origin of the almost ubiquitous presence of Arcobacter on poultry carcasses is still under discussion as the transmission routes of these bacteria are still not established. In contrast to the related Campylobacter, for which the contamination at broiler house level is well documented and easily detected by conventional microbiological methods, Arcobacter seem however to display a different behavior. Several authors have suggested that Arcobacter are probably not normal inhabitants of the poultry intestine and, as formulated by Eifert et al.,108 Houf et al.,113 Gude et al.,109 and Van Driessche and Houf94 that process water may be a potential source of the carcass contamination. An explanation for the contradictory reports in literature may be the sampling procedure. As in many studies, Arcobacter were isolated from the crates to transport the chickens to the slaughterhouse, one should take into account that those Arcobacter may contaminate the cloacal region, and may explain the isolations reported by some authors.37,98 As demonstrated in the study of Van Driessche and Houf,94 also the time and the sampling procedure are crucial and can affect the outcome of the study. Besides the reports of Arcobacter in chickens and turkeys, also the presence of Arcobacter in ducks and geese has been described.37,114–116 21.1.5.4  Arcobacter in Other Animals The occurrence of Arcobacter in horses has been examined by Van Driessche at al.26 who reported the isolation of A. butzleri out of two of the 15 examined animals. No information is available about the natural Arcobacter distribution in rodents, and a single report mentioned an A. cryaerophilus isolation from a naturally infected rainbow trout (Oncorhynchus mykiss Walbaum).70 The presence of Arcobacter spp. in raccoons

Number of Animals

Prevalence

250 50 50 10 20 55 20 60 14 250 78 294 144 134 100 72

5 0 6 20 5 36.3 0 0 7.1 10 44.8 41.1 42.3 59.7 39 55.6

Reference 106 96

98 26,92 105

(Procyon lotor) was reported for the first time in 2004.117 From a public health perspective, this observation may be important to note, since they share the immediate environment of human beings in certain countries and thus may play a part in the epidemiology of zoonotic bacterial infections. Arcobacter have been isolated both from ill and healthy nonhuman primates. Several cases have been reported of the isolation of A. butzleri from healthy infant macaques (Macaca nemestrina) in a monkey nursery facility, and from Rhesus macaque (Macaca mulatta) with and without diarrhea.29–31 The significance of these findings and whether they can serve as a model for human infection has not yet been determined.

21.1.6  Arcobacter in Food of Animal Origin Beside contaminated water, food of animal origin is another possible route of transmission of Arcobacter to humans. The exact routes of infection are not clear, but probably include manipulation of raw meat, the consumption of undercooked products and cross-contamination. Arcobacter, like thermotolerant campylobacters, have been reported more frequently from poultry products than from red meats. Recent studies have indicated that also Arcobacter are common on broiler carcasses. Arcobacter have also been isolated from skin samples of commercially reared ducks and turkeys. Eggs seem not to be infected. Apart from chickens, Arcobacter have been isolated from geese and ducks. A survey of mechanically separated turkey samples showed that this meat-type can be heavy contaminated with Arcobacter. A partial overview of the occurrence of Arcobacter on food of animal origin in different countries by multiple isolation protocols is shown in Table 21.3. At present no standard isolation method for Arcobacter has been proposed, therefore the true occurrence of Arcobacter, their contamination

295

Arcobacter

Table 21.3 Presence of Arcobacter on Food of Animal Origin Food Product Chicken carcass

Number of Samples

Prevalence

201

97

France

170

81

Germany

50

Country

Reference 119 82

52.3

USA

480

83

Belgium

120

75

95

Turkey

121

61

65.3

Czech Republic

122

22

73

Australia

123

41

48

Japan

124

10

100

Thailand

124

80

65

France

125

24.1

The Netherlands

52

65.4

Belgium

15

20

USA

100

23

Japan

94

62

Northern Ireland

127

57

0

Italy

128

395

77

USA

119

17

24

Denmark

27

Chicken meat 220

53 126 75 80

Eggs Turkey meat 37

Duck carcass 10

80

UK

10

70

Denmark

8

0

115 37

Rabbit meat Czech republic

122

Ground beef 45

28.9

USA

32

22

Australia

75

90

2,2

Japan

108

34

Northern Ireland

127

123 80

Minced beef 68

1.5

The Netherlands

53

97

5.1

Turkey

99

299

55.8

Ground pork 27

129

Italy

128

200

32

USA

130

21

29

Australia

123

100

7

101

35

Northern-Ireland

127

23.8

Belgium

131

21

3.7

USA

Japan

80

Minced pork 194

0.5

26

19.2

Belgium

The Netherlands

131

53

13

15

Australia

123

Sheep meat

296

level and their genotypic heterogeneity is largely unknown and limits the ability to compare field data. Furthermore, the variations in recovery rates can also be due to differences in country, farm management, hygiene in slaughterhouses and processing companies.118

21.1.7 Identification and Characterization of Arcobacter Members of the genus Arcobacter are Gram-negative nonspore forming organisms. Cells are usually slender, curved rods, 0.2–0.9 µm wide and 0.5–3 µm long. S-shaped or helical cells are often present. Cells in old cultures may form spherical or coccoid bodies and loose spiral filaments up to 20 µm long.132 Arcobacter are motile with a characteristic corkscrew-like motion by means of a single polar unsheathed flagellum at one or both ends of the cell.13 Although the cell surface is critical in pathogenic processes such as the colonization of the host, resistance to host defense systems and invasion of cells, the cell surface characteristics of Arcobacter are still largely unknown.133 Research about the presence of filamentous appendages, or specific proteins with porin and adhesive properties, which have received detailed research because of their role in the pathogenesis in both Campylobacter and Helicobacter species, has not been performed. Neither specific proteins in the S-layer, as described for Campylobacter fetus, nor polysaccharide components responsible for serotype specificity, are known to date. Optimal growth occurs at 30°C under microaerobic conditions, with a respiratory type of metabolism. Hydrogen is not required. After primary isolation in a microaerobic environment, growth is possible in aerobic or anaerobic atmosphere. Growth can occur at 15–37°C and growth at 42°C is described for some A. butzleri and A. skirrowii strains.134 Colonies of A. butzleri and A. cryaerophilus grown for 48 h under optimal conditions are respectively, 2–4 mm and 1–3 mm in diameter and are convex with an entire edge. Growth of A. butzleri has a whitish appearance whereas colonies of A. cryaerophilus have mostly a dirty yellow pigment. Colonies of A. skirrowii grown for 48 h under optimal conditions are 1–3 mm in diameter and have a flat irregular shape. Growth has a grayish appearance and is usually not profuse. Identification of Arcobacter at species level by biochemical characteristics is difficult as members of the genus display little metabolic activity.13,134 Classical biochemical differentiation of the genus Arcobacter from the related genera Campylobacter and Helicobacter, is primarily achieved by the identification of the individual species. In general, Arcobacter can be differentiated from Campylobacter by their lower optimal growth temperatures (25–30°C compared to 30–42°C for Campylobacter) and aerotolerance. However, in the identification of Campylobacteraceae at species level by the use of phenotypic tests, some fundamental problems can occur. First, many phenotypic tests used to differentiate other bacterial groups such as the members of the family Enterobacteriaceae have no discriminatory power for Campylobacteraceae because of their fastidious

Molecular Detection of Foodborne Pathogens

growth requirements and their relative metabolic inertness. For example, Arcobacter, and Campylobacter species do not ferment nor oxidize carbohydrates. Second, there is a lack of standardization for the tests that are used. For example, the outcome of a given test may be influenced by the inoculum size, cultural age, and basal medium used. The third problem is the lack of objectivity in the schemes available. Most tables so far described are used by comparing the test results of the unknown with the phenotypic profiles of known taxa and more importance is often attached to a single test result that is considered as an essential character than to the remainder of the phenotype. In a comprehensive study in 1996, On et al.135 documented an identification scheme for Campylobacteraceae and Helicobacter species, by which most Campylobacteraceae can be identified accurately and objectively with phenotypic tests when probabilistic methods of data assessment are employed. In contrast to other organisms as Salmonella, Campylo­ bacter, and Listeria, serology is not used for Arcobacter identification as attempts to perform genus or species identification by specific antibody agglutination were not successful.136 The high antigenic heterogeneity within the Arcobacter species may be on the basis of this failure. Due to the rather disappointing results, serological identification was not further extended, and is not further used to date. Differentiation between Arcobacter and Campylobacter isolates is possible by the determination of the cellular fatty acid composition. Arcobacter species possess a unique combination of a tetradecenoic acid C14:1 and two isomers of C16:1,137 later identified as C14:1ω7-cis, C16:1ω7-cis (common in most bacteria) and C16:1ω7-trans (considered unique for Arcobacter).138,139 Within the genus Arcobacter, A. nitrofigilis, A. skirrowii and the two A. cryaerophilus subgroups were differentiated by gas chromatography of the cellular fatty acids, but it was not able to differentiate A. butzleri from A. cryaerophilus subgroup 2.13 To date, due to the rather complex analysis protocol and the availability of faster, less complex and more cheaper molecular based methods, identification based on fatty acid profiles is not commonly applied, though it usefulness has been demonstrated by Jelinek et al.140 Identification based on whole-cell proteins profiles obtained by SDS-PAGE, has been the gold standard method since the description of the genus Arcobacter. By SDS-PAGE, all known Arcobacter species can be identified including the differentiation between A. cryaerophilus subgroup 1 and 2. However, an enforced standardization of the protocol combined with a profile library of known and related species and genera is necessarily. As this method is rather time consuming, it can hardly been applied in routine analysis. 21.1.7.1  Molecular Identification Differentiating of Arcobacter species by using phenotypic tests might give erroneous results because of the shortage of clear-cut differentiating tests, a phenomenon which has also been observed in the closely related genus Campylobacter. Therefore, several DNA-based assays were developed for

Arcobacter

the identification of Arcobacter at genus and species level. Recently, a microarray technique and a real-time fluorescence resonance energy transfer PCR for the detection and identification of Arcobacter spp. were reported, but are not routinely applied yet.141,142 RFLP. Using whole-cell chromosomal digests by the restriction enzyme PvuII and hybridization with probes derived from the Escherichia coli 16S and 23S rRNA genes, Kiehlbauch et al.33,143 developed a DNA-based method to differentiate the genera Campylobacter, Arcobacter, Helicobacter, and Wolinella. Although it is not able to distinguish A. butzleri from A. skirrowii, the method can be used for the differentiation of A. cryaerophilus from the other Arcobacter taxa and for the two subgroups of the latter species.144 Sequence analysis of the conserved 16S rRNA gene of Arcobacter allowed Wesley et al.144 to design a genus-specific nucleic acid probe and a species-specific DNA probe for A. butzleri. Southern blot hybridization of PvuII digested DNA using the Arcobacter genus-specific probe or the A. butzlerispecific probe end-labeled with [γ-32P]ATP provided a reliable identification method for Arcobacter at genus level and for A. butzleri. PCR. Analysis of the ribosomal gene sequence has proven to be a valuable tool in the determination of phylogenetic relationships between prokaryotes.10 A high (>94%) 16S rRNA gene sequences similarity was detected among the published Arcobacter species type strains. On the other hand, similarity to other members of the epsilon Proteobacteria was low (<90%).18,144 Based on the knowledge of the Arcobacter nucleic acid composition of the 16S rRNA, a genus- and ­species-specific DNA-probe was developed for identification of Arcobacter and A. butzleri strains.144 In recent years, identification was done using rapid and specific PCR methods. The 16S and 23S rRNA of living organisms contain information that reflects the evolutionary relation of bacteria. Specific primers, derived from conserved rRNA gene sequences, can be used to amplify genus- or species-specific regions. Different Arcobacter genus- and species-specific PCR assays have been described in literature and were reliable in the identification of reference strains and field isolates. One of the first described DNA-based identification techniques included a genus- and species-specific PCR reaction developed by Bastyns et al.145 with five primers targeting the 23S rRNA. One PCR amplification was necessary to identify the genus Arcobacter, a second PCR reaction could differentiate A. butzleri from other species and a third amplification distinguished A. cryaerophilus from A. skirrowii. The disadvantage of this technique was the need of DNA amplifications at different annealing temperatures. Based on the sequence of the Arcobacter and A. butzleri-specific DNAprobes described by Wesley et al.144, two primer pairs were developed that could be used in an Arcobacter genus- and an A. butzleri-specific PCR reaction with different annealing temperature.146 When the species-specific primers were replaced by the species-specific primers of Bastyns et al.,145 the first Arcobacter multiplex-PCR (m-PCR) was created: the genus Arcobacter (1223-bp) and the species A. butzleri

297

(686-bp) were identified in one PCR amplification.147 The genus- and species-specific PCR assays described so far were not able to detect all known Arcobacter species in one PCR amplification reaction. Therefore, m-PCR systems, targeting the 16S and 23S rRNA genes, have been developed for the simultaneous identification of the different human-related Arcobacter spp. in one PCR amplification and in one PCR tube.126 By means of five primers, a PCR product of 401-bp was generated for A. butzleri, 257-bp for A. cryaerophilus and 641-bp for A. skirrowii. Those three species were also identified by the PCR assay developed by Kabeya et al.,148 but A. cryaerophilus subgroup 1 and 2 were also differentiated from each other. Some identification protocols used PCR as a part of the identification method. A PCR-hybridization protocol differentiated A. butzleri from the related C. jejuni, C. coli, C. lari and C. upsaliensis strains.149 The conserved glyA gene region of isolates was amplified during a PCR reaction followed by a hybridization reaction of the amplicons with species-­specific oligodeoxyribonucleotide probes. Another example is a culture-PCR method, used to detect Arcobacter on chicken meat.150 After enrichment of the meat sample, identification of Arcobacter in the medium was performed by a new developed genus-PCR assay that generated an amplicon of 181-bp for Arcobacter positive samples. RFLP-PCR. Combined use of PCR with RFLP was first described by Cardarelli-Leite et al.151 RFLP analysis of a PCR-amplified DNA fragment of the gene coding for 16S rRNA of Campylobacter, Helicobacter, Arcobacter, and Wolinella succinogenes, generated a 283-bp fragment from all species belonging to the examined genera. Initial restriction of the amplicon by DdeI, delivered a unique pattern for A. butzleri. Performing additional digestion using HpaII on the DdeI digested fragments, A. cryaerophilus, A. skirrowii and A. nitrofigilis can be distinguished as a single group from the Campylobacter and Helicobacter species, although further differentiation at species level is not possible. Hurtado and Owen152 performed a comparable study, in which amplicons ranging from 2.6 to 3.0 kb are generated by a PCR assay using primers in the conserved region of the 23S rRNA gene of Campylobacter and Arcobacter. Digesting this amplicons with HaeIII, CfoI, HpaII, and HinfI, species-specific patterns for A. butzleri and A. nitrofigilis and identical patterns for A. cryaerophilus and A. skirrowii can be obtained. In 1999, Marshall et al.153 combined a PCR assay with RFLP for the identification of Arcobacter at the species level. By amplifying a 1004-bp fragment using primers targeting a conserved region of the 16S rRNA gene, followed by restriction endonuclease digestion with DdeI and TaqI, species specific RFLP patterns were obtained for A. butzleri, A. cryaerophilus and A. skirrowii. AFLP. The genotyping AFLP technique may also be used for concurrent species identification of the family Campylobacteraceae, including the Arcobacter species.38,154 The species A. butzleri, A. cryaerophilus and A. skirrowii form well-distinguished clusters in the dendrogram obtained after numerical analysis of yielded patterns.

298

21.1.7.2  Molecular-Based Characterization The determination of the relation of isolates below species level has become very important for the identification of transmission routes and biological reservoirs in epidemiological studies. Phenotyping methods such as biotyping,155 serotyping,156 or comparisons of whole-cell proteins,13 are of limited use due to their low discriminatory power and the instability and low reproducibility of the phenotypic characteristics.134 The suitability of pulsed-field gel electrophoresis (PFGE) and random amplification of polymorphic DNA (RAPD) as characterization techniques described by Lior and Wang157 was confirmed for fecal or meat isolates in recent reports.96,123,125,158,159 The AFLP technique proved its qualities as genotyping method for all members of the Campylobacteraceae.15,38,154 Good distinguished clusters were obtained for A. butzleri, A. cryaerophilus and A. skirrowii, with profiles reproducing the clonal relation from the isolates within each species. The AFLP-method is a robust method with high discriminatory power, but nevertheless, it is a demanding technique and requires a large reference database.66 An enterobacterial repetitive intergenic consensus (ERIC) PCR was optimized for the characterization of A. butzleri, A. cryaerophilus and A. skirrowii strains in combination with a rapid DNA extraction method.159 The technique is described in detail in Section 21.2.3.2. Fingerprints generated with ERIC-PCR are complex enough to differentiate at strain and substrain level and have a good reproducibility.

21.2  Methods 21.2.1  Arcobacter Isolation The isolation of Arcobacter is generally performed by a selective enrichment step, followed by plating on a selective or nonselective agar plate and takes approximately 4–5 days. This selective isolation is achieved predominantly by the use of antimicrobial agents in the enrichment and plating media or by incubations below 30°C. The first Arcobacter were coincidentally isolated using the semi-solid Ellinghausen-McCullough-Johnson-Harris (EMJH) medium containing 5-fluorouracil as selective supplement, originally developed for the isolation of Leptospira.4 Since the description of the genus Arcobacter, different isolation methods and protocols have been used to examine the presence of Arcobacter in different matrices. The first attempts were modifications of methods developed for thermophilic campylobacters or Yersinia species,58,129 and applied commercially available selective supplements, such as cefo perazone, amphotericin B, teicoplanin (CAT),160 cefoperazone, charcoal, deoxycholate agar (CCDA)128 or cefsulodin, irgasan, novobiocin (CIN).129 Those methods are either time­consuming, do not support the growth of all Arcobacter species or do not sufficiently inhibit the accompanying flora.78 The first specific selective Arcobacter isolation medium developed for Arcobacter isolation from meat and meat products contained

Molecular Detection of Foodborne Pathogens

cefoperazone, trimethoprim, piperacillin, and cycloheximide in the selective supplement.53 The isolation protocol consists of an enrichment in Arcobacter Selective Broth followed by plating on a semi-solid Arcobacter Selective Medium. After incubation, plates are examined for the presence of a motility zone. Since antibiotic sensitivity tests revealed that the piperacillin and cefoperazone concentrations in the medium inhibited the growth of A. skirrowii, this medium is not generally applied any longer. Another medium has been developed by Johnson and Murano in which the inhibition of the accompanying flora was obtained with a selective broth and agar containing cefoperazone and 5-fluorouracil as selective additives.161 A study on the MICs of selective agents on the growth of Arcobacter illustrated that most of the isolation media contained concentrations that inhibited the recovery of one or more Arcobacter species or did not sufficiently suppress the accompanying flora present in the sample.78 Based on these findings, a qualitative and quantitative isolation medium was developed by Houf et al.162 A broth and agar medium contained a combination of different antimicrobial agents as selective supplement: amphotericin B, cefoperazone, 5-­fluorouracil, novobiocin, and trimethoprim. The qualitative isolation procedure included enrichment in broth followed by plating on selective agar. Enumeration of the Arcobacter in the sample was performed by direct plating of 100 µl homogenate onto the selective agar plate. After microaerobic incubation, colonies were counted and the bacterial load could be determined. This protocol provides a fast and reliable method for the isolation and enumeration of all food-related Arcobacter species with a good suppression of the contaminating flora.

21.2.2  Sample Preparation Arcobacter can be detected directly in the enrichment media of food samples, in blood or fecal material or after isolation on selective or nonselective agar plate. Detection at enrichment phase or directly in the organic specimen is however often hindered by the presence of inhibiting components as blood, bile salts and organic material, in particular fat. Furthermore, a minimum number of Arcobacter must be present. The detection can however be enhanced by centrifugation of the enrichment broths or homogenates followed by several wash steps of the bacterial pellet to dilute the inhibiting components. Extraction of the DNA can then be performed by several commercially available DNA extraction kits, often designed specifically for a particular clinical specimen, or by heating the washed bacterial pellet for 15 min at 95°C or by a more extended DNA-extraction procedure as the guanidine thiocyanate extraction procedure according to Pitcher et al.163 described in detail below. The use of commercial extraction kits or the guanidine thiocyanate extraction procedure often results in more pure, nonfragmented DNA and is, after standardization of the amount of dsDNA, recommended for the molecular based characterization of Arcobacter isolates.

299

Arcobacter

Reagents RS buffer (pH 8) 0.1 M EDTA (ethylene diaminetetraactetic acid) 0.15 M NaCl – TE buffer (10 mM Tris-HCl pH 8, 1 mM EDTA) – GES solution (guanidium thiocyanate-EDTA-Sarkosyl) For 100 ml: 60 g guanidiumthiocyanate 20 ml 0.5 M EDTA 20 ml Milli Q 1 g N-lauroylsarcosine – NH4Ac (7.5 M) – Chloroform/isoamyl alcohol (24:1) – Isopropanol – Ethanol 70% DNA extraction protocol (1) Suspend an Arcobacter colony in 500 µl RS buffer (2) Centrifuge for 2 min at 15,000 × g and remove the supernatant (3) Wash the pellet in 500 µl RS buffer (4) Centrifuge 2 min at 15,000 × g and remove the supernatant (5) Suspend the pellet in 100 µl TE buffer (6) Add 500 µl GES solution and mix until cell lysis occur (7) Incubate for 10 min on ice (8) Add 250 µl cold NH4Ac and mix (9) Incubate for 10 min on ice (10) Add 500 µl cold chloroform/isoamylalcohol, and vortex (11) Centrifuge 20 min at 11,400 × g at 4°C (12) Collect as much as possible the upper layer without disturbing the visible interface (13) Add approximately 800 µl cold isopropanol and mix until strands of DNA are visible (14) Centrifuge 10 min at 15,000 × g and discard the supernatant (15) Wash the pellet three times with 100 µl 70% ethanol (16) Dry the pellet in air and dissolve in 100 µl TE buffer (17) Store at 4° or –20°C

21.2.3  Detection Procedures 21.2.3.1  Genus-specific Identification by PCR For a fast and reliable identification of Arcobacter at genus level, the PCR assay developed in 1996 by Harmon and Wesley146 can be applied. The assay utilizes primers ARCOI and ARCO II targeted to the genes encoding 16S rRNA, and a specific 1223-bp fragment is yielded for all presently described Arcobacter species. DNA may be extracted by commercially available DNA extraction kits, as well as by the guanidine thiocyanate-based method, or by simply boiling the isolates for 15 min at 95°C. After cooling and a short spin to pellet the cell debris, the whole-cell lysate is ready to

use in the assay. Quantification and standardization of the DNA extract or lysate is not necessary but should not exceed 200 ng dsDNA/µl. PCR protocol (1) Prepare the PCR mixture (50 µl) containing: – 1 × PCR buffer (10 mM Tris-HCl, 50 mM KCl) – 1.5 mM MgCl2 – Primers: 50 pmol of ARCO I: 5′-AGA GAT TAG CCT GTA TTG TAT-3′ ARCO II: 5 ′-TAG CAT CCC CGC TTC GAA TGA-3′ – 200 µM dATP, dTTP, dGTP, and dCTP – 1.25 U Taq DNA polymerase – 1–2 µl of DNA extract or lysate (2) Run the following cycling program on a thermal cycler: one cycle of 94°C for 4 min; 25 cycles of 94°C for 1 min, 56°C for 1 min, and 72°C for 1 min. (3) Separate the amplified products on 1.5% agarose gel (in 1 × TAE or 0.5 × TBE buffer) containing 0.5 µg/ml of ethidium bromide and visualize on an UV transilluminator. 21.2.3.2  Species-specific Identification by PCR A species-specific PCR assay for the identification of A. butzleri, A. cryaerophilus 1A and 1B and A. skirrowii has been developed by Kabeya et al.148 The species-­specific forward primers from the most variable region of the 23S rRNA gene are N.butz, N.c1.A, N.c.1B, and N.ski, and the reverse primer ARCO-U recognizes a conserved Arcobacter specific region. DNA may be extracted by commercially available DNA extraction kits, as well as by the guanidium thiocyanate-based method, or by simply boiling the isolates for 15 min at 95°C. After cooling and a short spin to pellet the cell debris, the whole-cell lysate is ready to use in the assay. Authors recommend adjusting the DNA concentration to 10 ng DNA/µl. In the one-step PCR assay a specific 728-bp fragment is generated for A. cryaerophilus 1A, a 692-bp fragment for A. butzleri, a 488-bp fragment for A. skirrowii and a 152-bp fragment for A. cryaerophilus 1B. PCR protocol (1) Prepare PCR mixture (20 µl) containing: – 1 × PCR buffer (10 mM Tris-HCl, 50 mM KCl) – 1.5 mM MgCl2 – Primers: 50 pmol of N.butz 5′-AGCGTTCTAT TCAGCGTAGAAGATGT-3′ N.c1A 5′-ACCGAAGCTTTAGATTCGAATT TATTCG-3′ N.c1B 5′-GGACTTGCTCCAAAAAGCTGA AG-3′ N.ski 5′-CGAGGTCACGGATGGAAGTG-3′ ARCO-U 5′-TTCGCTTGCGCTGACAT CAT-3′ – 200 µM dATP, dTTP, dGTP, and dCTP

300

– 1.0 U Taq DNA polymerase – 2 µl of DNA extract or lysate (2) Perform PCR amplification as follows: one cycle of 94°C for 3 min; 30 cycles of 94°C for 30 sec, 62°C for 1 min and 72°C for 1 min; one cycle of 72°C for 5 min. (3) Separate the amplified products on 2% agarose gel (in 1 × TAE or 0.5 × TBE) containing 0.5 µg/ml of ethidium bromide and visualize on an UV trans­ illuminator. 21.2.3.3  Species Identification by PCR-RFLP As none of the above mentioned methods are able to discriminate the six currently accepted species, a 16S rDNARFLP method for the identification of all Arcobacter species was developed by Figueras et al.164 The assay comprises an initial amplification of a 1026-bp fragment targeted the 16S rRNA gene using a modified PCR assay of Marchall et al.153 followed by an endonuclease digestion and electrophoresis of the fragments. DNA may be extracted by commercially available DNA extraction kit, by a guanidine thiocyanate- or phenol/chloroform-based method, or by preparing a wholecell lysate by boiling. The DNA concentration should be adjusted to 100 ng dsDNA/µl. PCR protocol (1) Prepare PCR mixture (50 µl) containing: - PCR buffer (10 mM Tris-HCL, 50 mM KCl) - 1.5 mM MgCl2 - Primers: 50 pmol of mCAH1a 5′-AAC ACA TGC AAG TCG AAC GA-3′ CAH1b 5 ′-TTA ACC CAA CAT CTC ACG AC-3′ - 200 µM dATP, dTTP, dGTP, and dCTP - 2.5 U Taq DNA polymerase - 5 µl whole-cell lysate (2) Perform PCR amplification using following cycling program: one cycle of 95°C for 2 min; 30 cycles of 94°C for 30 sec, 52°C for 30 sec and 72°C for 90 sec; and one cycle of 72°C for 5 min. (3) Digest 10 μl of the amplification product with 10 U of the enzyme MseI (and 2 μl of the 10 × buffer R) in a total volume of 20 μl at 65°C for 3 h. (4) Separate the restriction fragments on 15% polyacrylamide gel electrophoresis in 1 × TBE buffer at 350 V for 5 h. (5) Stain the gel with ethidium bromide and photograph on a UV transilluminator. 21.2.3.4  Characterization by ERIC-PCR Houf et al. optimized an ERIC-PCR for the genetic characterization of Arcobacter at strain level and assessed the performance and discriminatory abilities of this method.159 In this and later studies, ERIC-PCR has shown to be a valuable, fast, and robust technique for characterizing A. butzleri, A. cryaerophilus, A. skirrowii and A. cibarius isolates.18,93,159

Molecular Detection of Foodborne Pathogens

PCR protocol (1) Prepare PCR mixture (50 µl) containing: – PCR buffer (10 mM Tris-HCL, 50 mM KCl) – 4 mM MgCl2 – primers: 25 pmol of ERIC1R: 5′-ATGTAAGC TCCTGGGGATTCAC-3′ ERIC2: 5 ′-AAGTAAGTGACTGGGGTGAG CG-3′ – 200 µM dATP, dTTP, dGTP, and dCTP – 5 U Taq DNA polymerase – 1 µl of DNA extract or 1 µl whole-cell lysate (2) Perform PCR amplification as follows: one cycle of 94°C for 5 min; 40 cycles of 94°C for 1 min, 25°C for 1 min, and 72°C for 2 min. (3) Separate the PCR products (8 µl) on 2% agarose gel (in 0.5 × TBE) containing 0.5 µg/ml ethidium bromide for 2.5 h at 100 V. Note: the banding patterns used to determine the ERIC-PCR type comprise DNA fragments between 100 and 2072 bp. For the interpretation of the fingerprints, use computer-based normalization and interpolation of the DNA profiles, and numerical analysis using the Pearson product moment correlation coefficient, with 1% position tolerance. Construct dendrograms using the unweighted pair group linkage analysis method (UPGMA). DNA patterns that differ in one or more DNA-fragments represent different types. Furthermore, groups of identical fingerprints, or fingerprints with slight differences in band intensity, are characterized by Pearson correlation coefficients of 90% or higher.

21.3 Conclusions and Future Perspectives Arcobacter are commonly isolated from animals, food, and environmental samples worldwide, and cases of human Arcobacter infection have regularly been reported. The infection source remains however often unclear. Several major aspects actually hamper the risk assessment for these bacteria. First, besides the fact that clinical samples are not routinely tested for Arcobacter species as is done for Salmonella or Campylobacter, most isolations are performed by methods designed for thermophilic Campylobacter species. The selectivity of those media is mostly achieved by the incorporation of antimicrobial agents and though Arcobacter have sporadically been isolated in this way, several studies about the Arcobacter susceptibility to the antimicrobials included showed that none of those supplements allowed the growth of all Arcobacter species or strains and at the same time sufficiently suppressed the accompanying flora present in biological samples. The low recovery rate reported today from human diarrheal specimens is therefore certainly an underestimation of the real prevalence. In order to determine the exposure assessment of Arcobacter to humans, efficient, robust, and reliable methods for isolation, identification, and characterization of Arcobacter are needed. Validation should

301

Arcobacter

be based on the determination of the parameters specificity, sensitivity, repeatability, reproducibility, and detection limit, both for classical bacteriological isolation as for molecularbased methods. Correct identification of Arcobacter is another challenge in microbiology, and too often, Arcobacter are misidentified as campylobacters, especially when phenotypical methods are applied. Due to the relatively metabolic inertness and the antigenic heterogeneity, biochemical or serological identification of Arcobacter is not recommended. Besides, they are time consuming and laborious and clear-cut parameters are not always present in all strains of a certain species. For now, identification at genus level can reliable be performed by genus-specific PCR assays, but species identification is often limited to some of the well-established species. For the more recently described species as A. cibarius and potential new species yet to be described, no or not routinely applicable methods are available. Furthermore, as most of the one-step PCR assays target the 16S or 23S genes, it can be expected that some new species can not be distinguished by this approach as only few differences exist between the several species at gene level. Though the PCR-RFLP approach has shown to be the only routinely applicable method to differentiate the six species at present, the occurrence of single mutations or the presence of the same cleaving place within two species cannot be excluded. In the future, genome analysis will probably identify more stable and discriminatory genes to create a more robust identification assay. Direct detection of Arcobacter by PCR, or even direct quantification by RT-PCR in a biological specimen is not commonly applied at this moment. In contrast to more vulnerable bacteria as Helicobacter species, Arcobacter are relatively easily to culture and there is no demand in human or veterinary medicine for a rapid analysis tool at the moment. Furthermore, as biological matrices are complex and often comprise inhibiting factors, preparation, and clean-up of the sample is expensive and hamper their use in routine laboratories. Furthermore, many studies are focused on further examination of the isolates as microbiological susceptibility and typing for which the preservation of the isolates is needed. The use of those direct methods can however have a future in large scale surveys. The existence of considerable heterogeneity among Arco­ bacter in one specimen is yet another difficulty in Arcobacter research. This phenomenon was already reported in early biotyping and serotyping studies. Possible explanations are multiple sources of contamination, the existence of multiple parent genotypes, and a high degree of genomic recombination among the progeny of parent genotypes, but none of these hypotheses has been demonstrated yet. Furthermore, in vitro cultivation of Arcobacter isolates over a period of time did not induce genomic rearrangements as identical fingerprints were obtained in the reproducibility studies. The use of molecular typing has also shown that this heterogeneity is not linked to artefacts by phenotypic characterization, but truly exists in the Arcobacter population. As a result, typing Arcobacter isolates often yields an

overwhelming jumble of different genotypes with no extra contribution to the ­identification of possible transmission routes. Also a bias caused by the isolation methods has clearly been demonstrated. Therefore, although commonly applied in current food and medical molecular microbiology, typing at strain level for such heterogeneous bacteria should seriously be considered and aims clearly stated before outset. Typing in the first place should be reserved to determine the relation between a direct potential contamination source and the patient’s isolate for which also epidemiological based evidence is available. When typing is appropriate, several methods are available, though only molecular based systems are currently applied. In practice, the choice often depends on the availability of equipment, knowledge, and experience in the lab. Several studies have shown that ERIC-PCR or RAPD are easy to perform, and have shown to be reproducible in their outcome. For fast, cheap, and easy to perform characterization needs, ERICPCR is therefore recommended as first line tool, but can be complemented by AFLP, MLST or by a non-PCR based method as PFGE. However, the latter methods are far more demanding and do not necessarily have a larger discriminatory power. Standardization of the DNA amount analysed in a PCR assay is often ­recommended, but the use of a DNA extraction by commercial kits, in house extraction methods or a simply boiling of the isolates to ­prepare whole-cell extracts as such result in the same outcome of the genotypic relatedness of the isolates. However, as they have an influence of the fingerprint patterns, they can not be combined within the same analysis. Finally, to establish epidemiological links between Arcobacter isolated from different to closely related sources or to elucidate the transmission routes, the clonally relatedness and the significance of a single fragment difference as often noticed in PCR-based fingerprints should be elucidated. The application of MLST and the set up of a MLST-type bank, as established for the closely related Campylobacters, may be helpful to accomplish these goals.

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Arcobacter 49. Romero, J. et al. Bacterial 16S rRNA gene analysis revealed that bacteria related to Arcobacter spp. constitute an abundant and common component of the oyster microbiota (Tiostrea chilensis). Microbial. Ecol., 44, 365, 2002. 50. Wirsen, C.O. et al. Characterization of an autotrophic ­sulfide-oxidizing marine Arcobacter sp that produces filamentous sulfur. Appl. Environ. Microbiol., 68, 316, 2002. 51. Hammes, F. et al. Molecular, biochemical and ecological characterisation of a bio-catalytic calcification reactor. Appl. Environ. Microbiol., 62, 191, 2003. 52. Grabowski, A. et al. Microbial diversity in production waters of a low-temperature biodegraded oil reservoir. FEMS Microbiol. Ecol., 54, 427, 2005. 53. de Boer, E. et al. A selective medium for the isolation of Arcobacter from meats. Lett. Appl. Microbiol., 23, 64, 1996. 54. Engberg, J. et al. Prevalence of Campylobacter, Arcobacter, Helicobacter, and Sutterella spp. in human fecal samples as estimated by a reevaluation of isolation methods for campylobacters. J. Clin. Microbiol., 38, 286, 2000. 55. Lastovica, A.J. and le Roux, E. Efficient isolation of campylobacteria from stools. J. Clin. Microbiol., 38, 2798, 2000. 56. Tee, W. et al. Campylobacter cryaerophila isolated from a human. J. Clin. Microbiol., 26, 2469, 1988. 57. Lerner, J. et al. Severe diarrhea associated with Arcobacter butzleri. Eur. J. Clin. Microbiol. Infect. Dis., 13, 660, 1994. 58. Burnens, A.P. et al. Isolation of Arcobacter butzleri from a girl with gastroenteritis on Yersinia selective CIN agar. Med. Microbiol. Lett., 251, 1992. 59. Lau, S.K.P. et al. Identification by 16S ribosomal RNA gene sequencing of Arcobacter butzleri bacteraemia in a patient with acute gangrenous appendicitis. Mol. Pathol., 55, 182, 2002. 60. Vandamme, P. et al. Outbreak of recurrent abdominal cramps associated with Arcobacter butzleri in an Italian school. J. Clin. Microbiol., 30, 2335, 1992. 61. Rice, E.W. et al. Isolation of Arcobacter butzleri from ground water. Lett. Appl. Microbiol., 28, 31, 1999. 62. On, S.L.W. et al. Isolation of Arcobacter butzleri from a neonate with bacteremia. J. Infect., 31, 225, 1995. 63. Yan, J.J. et al. Arcobacter butzleri bacteremia in a patient with liver cirrhosis. J. Formos. Med. Assoc., 99, 166, 2000. 64. Hsueh, P.R. et al. Bacteremia caused by Arcobacter cryaerophilus 1B. J. Clin. Microbiol., 35, 489, 1997. 65. Woo, P.C.Y. et al. Identification of Arcobacter cryaerophilus isolated from a traffic accident victim with bacteremia by 16S ribosomal RNA gene sequencing. Diagn. Microbiol. Infect. Dis., 40, 125, 2001. 66. Phillips, C.A. Arcobacter spp in food: isolation, identification and control. Trends Food Sci. Technol., 12, 263, 2001. 67. Ho, H.T.K. et al. Arcobacter, what is known and unknown about a potential foodborne zoonotic agent!, Vet. Microbiol., 115, 1, 2006. 68. Johnson, L.G. and Murano, E.A. Lack of a cytolethal distending toxin among Arcobacter isolates from varioussources. J. Food Prot., 65, 1789, 2002. 69. Wesley, I.V. et al. Infection of cesarean-derived ­colostrum-deprived 1-day-old piglets with Arcobacter butzleri, Arcobacter cryaerophilus, and Arcobacter skirrowii. Infect. Immun., 64, 2295, 1996. 70. Yildiz, H. and Aydin, S. Pathological effects of Arcobacter cryaerophilus infection in rainbow trout (Oncorhynchus mykiss Walbaum). Acta Veterinaria Hungarica, 54, 191, 2006.

303 71. Tsang, R.S.W. et al. Immunochemical characterization of a haemagglutinating antigen of Arcobacter spp. FEMS Microbiol. Lett., 136, 209, 1996. 72. Carbone, M. et al. Adherence of environmental Arcobacter butzleri and Vibrio spp. isolates to epithelial cells in vitro. Food Microbiol., 20, 611, 2003. 73. Fernandez, H. et al. Toxigenic and invasive capacities: possible pathogenic mechanisms in Arcobacter cryaerophilus. Memorias do Instituto Oswaldo Cruz, 90, 633, 1995. 74. Kalman, M., Czermann, B. and Szöllosy, E. Arcobacter butzleri strains in Csongrad county and their cytotoxin production in different cell lines. Acta Microbiologica et Immunologica Hungarica, 143, 1996. 75. Villarruel-Lopez, A. et al. Isolation of Arcobacter spp. from retail meats and cytotoxic effects of isolates against Vero cells. J. Food Prot., 66, 1374, 2003. 76. Carter, E.R. Enteropathogenicity of Arcobacter butzleri in rabbit and pig ileal loops. Thesis Master of Science, Faculty of the Virginia Polytechnic Institute and State University, Blacksburg, VA, 1996. 77. Fera, M.T. et al. In vitro susceptibility of Arcobacter butzleri and Arcobacter cryaerophilus to different antimicrobial agents. Int. J. Antimicrob. Agents, 21, 488, 2003. 78. Houf, K. et al. Susceptibility of Arcobacter butzleri, Arcobacter cryaerophilus, and Arcobacter skirrowii to antimicrobial agents used in selective media. J. Clin. Microbiol., 39, 1654, 2001. 79. Kiehlbauch, J.A. et al. In vitro susceptibilities of aerotolerant Campylobacter isolates to 22 antimicrobial agents. Antimicrob. Agents Chemother., 36, 717, 1992. 80. Kabeya, H. et al. Prevalence of Arcobacter species in retail meats and antimicrobial susceptibility of the isolates in Japan. Int. J. Food Microbiol., 90, 303, 2004. 81. Atabay, H.I. and Aydin, F. Susceptibility of Arcobacter butzleri isolates to 23 antimicrobial agents. Lett. Appl. Microbiol., 33, 430, 2001. 82. Harrass, B. et al. Identification and characterization of Arcobacter isolates from broilers by biochemical tests, antimicrobial resistance patterns and plasmid analysis. Zentralbl. Veterinarmed. B, 45, 87, 1998. 83. Vandenberg, O. et al. Antimicrobial susceptibility of clinical isolates of non-jejuni/coli campylobacters and arcobacters from Belgium. J. Antimicrob. Chemother., 57, 908, 2006. 84. Hakanen, A. et al. Quality control strains used in susceptibility testing of Campylobacter spp. J. Clin. Microbiol., 40, 2705, 2002. 85. Luber, P. et al. Comparison of broth microdilution, E test, and agar dilution methods for antibiotic susceptibility testing of Campylobacter jejuni and Campylobacter coli. J. Clin. Microbiol., 41, 1062, 2003. 86. Houf, K. et al. Antimicrobial susceptibility patterns of Arcobacter butzleri and Arcobacter cryaerophilus strains isolated from humans and broilers. Microb. Drug Resist. 10, 243, 2004. 87. Schroeder-Tucker, L. et al. Phenotypic and ribosomal RNA characterization of Arcobacter species isolated from porcine aborted fetuses. J. Vet. Diagn. Invest., 8, 186, 1996. 88. On, S.L.W. et al. Prevalence and diversity of Arcobacter spp. isolated from the internal organs of spontaneous porcine abortions in Denmark. Vet. Microbiol., 85, 159, 2002. 89. de Oliveira, S.J. et al. Classification of Arcobacter species isolated from aborted pig fetuses and sows with reproductive problems in Brazil. Vet. Microbiol., 57, 347, 1997.

304 90. Logan, E.F. et al. Mastitis in dairy cows associated with an aerotolerant Campylobacter. Vet. Rec., 110, 229, 1982. 91. Suarez, D.L. et al. Detection of Arcobacter species in gastric samples from swine. Vet. Microbiol., 57, 325, 1997. 92. Van Driessche, E. et al. Occurrence and strain diversity of Arcobacter species isolated from healthy Belgian pigs. Res. Microbiol., 155, 662, 2004. 93. Van Driessche, E. et al. Prevalence, enumeration and strain variation of Arcobacter species in the faeces of healthy cattle in Belgium. Vet. Microbiol., 105, 149, 2005. 94. Van Driessche, E. and Houf, K. Discrepancy between the occurrence of Arcobacter in chickens and broiler carcass contamination. Poult. Sci., 86, 744, 2007. 95. Wesley, I.V. et al. Fecal shedding of Campylobacter and Arcobacter spp. in dairy cattle. Appl. Environ. Microbiol., 66, 1994, 2000. 96. Hume, M.E. et al. Genotypic variation among Arcobacter isolates from a farrow-to-finish swine facility. J. Food Prot., 64, 645, 2001. 97. Golla, S.C. et al. Determination of the occurrence of Arcobacter butzleri in beef and dairy cattle from Texas by various isolation methods. J. Food Prot., 65, 1849, 2002. 98. Kabeya, H. et al., Distribution of Arcobacter species among livestock in Japan. Vet. Microbiol., 93, 153, 2003. 99. Ongor, H. et al. Investigation of arcobacters in meat and faecal samples of clinically healthy cattle in Turkey. Lett. Appl. Microbiol., 38, 339, 2004. 100. Fernandez, H. et al. First isolation in Chile of Arcobacter cryaerophilus from a bovine abortion. Archivos de Medicina Veterinaria, 27, 111, 1995. 101. Parvanta, M.F. Campylobacter cryaerophila and Campylobacter fetus subspecies venerealis as a cause of serial abortions in two cattle herds in North-Rhine-Westfalia. Tierarztliche Umschau, 54, 364, 1999. 102. Gill, K.P. Aerotolerant campylobacter strain isolated from a bovine preputial sheath washing. Vet. Rec., 112, 459, 1983. 103. de Oliveira, S.J. et al. Antigenic diversity among strains of Arcobacter spp. isolated from pigs in Rio Grande do Sul, Brazil and presence of agglutinin titers in serum samples of sows with reproductive problems. Ciência Rural, Santa Maria, 29, 705, 1999. 104. Jahn, B. Campylobacter im genitaltrakt des schweines. Inaugural dissertation, Tierärztliche Hochshule Hannover, Hannover, Germany, 1993. 105. Ho, T.K.H. et al. Potential routes of acquisition of Arcobacter species by piglets. Vet. Microbiol., 114, 123, 2006. 106. Harvey, R.B. et al. Prevalence of Campylobacter, Salmonella, and Arcobacter species at slaughter in market age pigs. Adv. Exp. Med. Biol., 473, 237, 1999. 107. Atabay, H.I. et al. Diversity and prevalence of Arcobacter spp. in broiler chickens. J. Appl. Microbiol., 84, 1007, 1998. 108. Eifert, J.D. et al. Comparison of sampling techniques for detection of Arcobacter butzleri from chickens. Poult. Sci., 82, 1898, 2003. 109. Gude, A. et al. Ecology of Arcobacter species in chicken rearing and processing. Lett. Appl. Microbiol., 41, 82, 2005. 110. Talhouk, R.S. et al. Prevalence, antimicrobial susceptibility and molecular characterization of Campylobacter isolates recovered from humans and poultry in Lebanon. J. Med. Liban., 46, 310, 1998. 111. Corry, J.E.L. and Atabay, H.I. Poultry as a source of Campylobacter and related organisms. J. Appl. Microbiol., 90, 96S, 2001.

Molecular Detection of Foodborne Pathogens 112. Wesley, I.V. and Baetz, A.L. Natural and ­experimental infections of Arcobacter in poultry. Poult. Sci., 78, 536, 1999. 113. Houf, K. et al. Molecular characterization of Arcobacter isolates collected in a poultry slaughterhouse. J. Food Prot., 66, 364, 2003. 114. Atabay, H.I. et al. Prevalence of Arcobacter species in domestic geese (Anser anser) in Turkey. Zoonoses Public Hlth., 54, 73, 2007. 115. Ridsdale, J.A. et al. Prevalence of campylobacters and arcobacters in ducks at the abattoir. J. Appl. Microbiol., 85, 567, 1998. 116. Ridsdale, J.A. et al. Campylobacter and Arcobacter spp. isolated from the carcasses and caeca of commercially reared ducks. Anaerobe, 5, 317, 1999. 117. Hamir, A.N. et al. Campylobacter jejuni and Arcobacter species associated with intussusception in a raccoon (Procyon lotor). Vet. Rec., 155, 338, 2004. 118. Phillips, C.A. Arcobacters as emerging human foodborne pathogens. Food Control, 12, 1, 2001. 119. Manke, T.R. et al. Prevalence and genetic variability of Arcobacter species in mechanically separated turkey. J. Food Prot., 61, 1623, 1998. 120. Johnson, L.G. and Murano, E.A. Comparison of three protocols for the isolation of Arcobacter from poultry. J. Food Prot., 62, 610, 1999. 121. Atabay, H.I. et al. The prevalence of Arcobacter spp. on chicken carcasses sold in retail markets in Turkey, and identification of the isolates using SDS-PAGE. Int. J. Food Microbiol., 81, 21, 2003. 122. Vytrasova, J. et al. Isolation of Arcobacter butzleri and A. cryaerophilus in samples of meats and from meat-processing plants by a culture technique and detection by PCR. Folia Microbiologica, 48, 227, 2003. 123. Rivas, L. et al. Isolation and characterisation of Arcobacter butzleri from meat. Int. J. Food Microbiol., 91, 31, 2004. 124. Morita, Y. et al. Isolation and phylogenetic analysis of Arcobacter spp. in ground chicken meat and environmental water in Japan and Thailand. Microbiol. Immun., 48, 527, 2004. 125. Festy, B., et al. Poultry meat and water as the possible sources of Arcobacter butzleri associated human disease in Paris, France. Acta-enterogastrologica Belgica, Suppl. 6, 35, 1993. 126. Houf, K. et al. Development of a multiplex PCR assay for the simultaneous detection and identification of Arcobacter butzleri, Arcobacter cryaerophilus and Arcobacter skirrowii. FEMS Microbiol. Lett., 193, 89, 2000. 127. Scullion, R. et al. Prevalence of Arcobacter spp. in raw milk and retail raw meats in northern Ireland. J. Food Prot., 69, 1986, 2006. 128. Zanetti, F. et al. Prevalence of thermophilic Campylobacter and Arcobacter butzleri in food of animal origin. Int. J. Food Microbiol., 33, 315, 1996. 129. Collins, C.I. et al. Detection of Arcobacter spp in ground pork by modified plating methods. J. Food Prot., 59, 448, 1996. 130. Ohlendorf, D.S. and Murano, E.A. Prevalence of Arcobacter spp. in raw ground pork from several geographical regions according to various isolation methods. J. Food Prot., 65, 1700, 2002. 131. Van Driessche E. and Houf, K. Characterization of the Arcobacter contamination on Belgian pork carcasses and raw retail pork. Int. J. Food Microbiol., 118, 20, 2007.

Arcobacter 132. Mansfield, L.P. and Forsythe, S.J. Arcobacter butzleri, A. skirrowii and A. cryaerophilus - potential emerging human pathogens. Rev. Med. Microbiol., 11, 161, 2000. 133. Penn, C.W. Campylobacter, Arcobacter and Helicobacter: surface characteristics. In: Campylobacter, Helicobacter and Arcobacter. Summer 2000 Conference, Society for Applied Microbiology, Strathclyde, UK, 3, 2000. 134. On, S.L.W. Identification methods for campylobacters, helicobacters, and related organisms. Clin. Microbiol. Rev., 9, 405, 1996. 135. On, S.L.W. et al. A probability matrix for the identification of campylobacters, helicobacters and allied taxa. J. Appl. Bacteriol., 81, 425, 1996. 136. Boudreau, M. et al. Biochemical and serological characterization of Campylobacter cryaerophila. J. Clin. Microbiol., 29, 54, 1991. 137. Lambert, M.A. et al. Differentiation of Campylobacter and Campylobacter-like organisms by cellular fatty acid composition. J. Clin. Microbiol., 25, 706, 1987. 138. Moss, C.W. and Lambert-Fair, M.A. Location of double bonds in monounsaturated fatty acids of Campylobacter cryaerophila with dimethyl disulfide derivatives and combined gas chromatography-mass spectrometry. J. Clin. Microbiol., 27, 1467, 1989. 139. Moss, C.W. et al. Isoprenoid quinones of Campylobacter cryaerophila, C. cinaedi, C. fennelliae, C. hyointestinalis, C. pylori, and “C. upsaliensis”. J. Clin. Microbiol., 28, 395, 1990. 140. Jelinek, D. et al. Identification of Arcobacter species using phospholipid and total fatty acid profiles. Folia Microbiologica, 51, 329, 2006. 141. Abdelbaqi, K. et al. Development of a real-time fluorescence resonance energy transfer PCR to detect Arcobacter species. J. Clin. Microbiol., 45, 3015, 2007. 142. Miller, W.G. et al. Novel multilocus sequence typing methods for multiple human pathogenic species of Arcobacter reveal substantial diversity within the genus. In: 107th General Meeting of the American Society for Microbiology, Metro Toronto Convention Centre, Toronto, Canada, C-305, 2007. 143. Kiehlbauch, J.A. et al. Evaluation of ribotyping techniques as applied to Arcobacter, Campylobacter and Helicobacter. Mol. Cell. Probes, 8, 109, 1994. 144. Wesley, I.V. et al. Arcobacter-specific and Arcobacter butzleri-specific 16S ribosomal-RNA-based DNA probes. J. Clin. Microbiol., 33, 1691, 1995. 145. Bastyns, K. et al. A variable 23S rDNA region is a useful discriminating target for genus-specific and species-specific PCR amplification in Arcobacter species. Syst. Appl. Microbiol., 18, 353, 1995. 146. Harmon, K.M. and Wesley, I.V. Identification of Arcobacter isolates by PCR. Lett. Appl. Microbiol., 23, 241, 1996. 147. Harmon, K.M. and Wesley, I.V. Multiplex PCR for the identification of Arcobacter and differentiation of Arcobacter butzleri from other arcobacters. Vet. Microbiol., 58, 215, 1997.

305 148. Kabeya, H. et al. One-step polymerase chain ­reaction-based typing of Arcobacter species. Int. J. Food Microbiol., 81, 163, 2003. 149. Al Rashid, S.T. et al. Identification of Campylobacter jejuni, C. coli, C. lari, C. upsaliensis, Arcobacter butzleri, and A. butzleri-like species based on the glyA gene. J. Clin. Microbiol., 38, 1488, 2000. 150. Gonzalez, I. et al. Development of a combined PCRculture technique for the rapid detection of Arcobacter spp. in chicken meat. Lett. Appl. Microbiol., 30, 207, 2000. 151. Cardarelli-Leite, P. et al. Rapid identification of Campylo­ bacter species by restriction fragment length polymorphism analysis of a PCR-amplified fragment of the gene coding for 16S rRNA. J. Clin. Microbiol., 34, 62, 1996. 152. Hurtado, A. and Owen, R.J. A molecular scheme based on 23S rRNA gene polymorphisms for rapid identification of Campylobacter and Arcobacter species. J. Clin. Microbiol., 35, 2401, 1997. 153. Marshall, S.M. et al. Rapid identification of Campylobacter, Arcobacter, and Helicobacter isolates by PCR-restriction fragment length polymorphism analysis of the 16S rRNA gene. J. Clin. Microbiol., 37, 4158, 1999. 154. On, S.L.W. et al. Genotyping and genetic diversity of Arcobacter butzleri by amplified fragment length polymorphism (AFLP) analysis. Lett. Appl. Microbiol., 39, 347, 2004. 155. Lior, H. and Woodward, D.L. Arcobacter butzleri: a biotyping scheme. Acta Gastro-enterologica Belgica, Suppl. 6, 28, 1993. 156. Lior, H. and Woodward, D.L. Arcobacter butzleri: a serotyping scheme. Acta Gastro-enterologica Belgica, Suppl. 6, 29, 1993. 157. Lior, H. and Wang, G. Differentiation of Arcobacter butzleri by pulsed field gel electrophoresis (PFGE) and random amplified polymorphic DNA (RAPD). Acta Gastroenterologica Belgica, Suppl. 6, 29, 1993. 158. Atabay, H.I. et al. Discrimination of Arcobacter butzleri isolates by polymerase chain reaction-mediated DNA fingerprinting. Lett. Appl. Microbiol., 35, 141, 2002. 159. Houf, K. et al. Assessment of the genetic diversity among arcobacters isolated from poultry products by using two PCR-based typing methods. Appl. Environ. Microbiol., 68, 2172, 2002. 160. Atabay, H.I. and Corry, J.E.L. The prevalence of campylobacters and arcobacters in broiler chickens. J. Appl. Microbiol., 83, 619, 1997. 161. Johnson, L.G. and Murano, E.A. Development of a new medium for the isolation of Arcobacter spp. J. Food Prot., 62, 456, 1999. 162. Houf, K. et al. Development of a new protocol for the isolation and quantification of Arcobacter species from poultry products. Int. J. Food Microbiol., 71, 189, 2001. 163. Pitcher, D.G. et al. Rapid extraction of bacterial genomic DNA with guanidium thiocyanate. Lett. Appl. Microbiol., 8, 151, 1989. 164. Figueras, M.J., Collado, L. and Guarro, J. A new 16S rDNA-RFLP method for the discrimination of the accepted species of Arcobacter. Diagn. Microbiol. Infect. Dis., 62, 11, 2008.

22 Bacteroides

Rama Chaudhry, Anubhav Pandey, and Nidhi Sharma All India Institute of Medical Sciences

Contents 22.1 Introduction.................................................................................................................................................................... 307 22.1.1 Biological Characteristics ................................................................................................................................ 307 22.1.2 Epidemiology.................................................................................................................................................... 308 22.1.3 Pathogenesis...................................................................................................................................................... 309 22.1.4 Enterotoxin: Virulence Factor of B. fragilis..................................................................................................... 309 22.1.5 Role of Molecular Methods for B. fragilis.........................................................................................................310 22.2 Methods...........................................................................................................................................................................311 22.2.1 Reagents and Equipment....................................................................................................................................311 22.2.2 Sample Collection and Preparation ..................................................................................................................311 22.2.3 Detection Procedures.........................................................................................................................................312 22.2.3.1 DNA Extraction................................................................................................................................312 22.2.3.2 PCR for Neuraminidase Gene .........................................................................................................314 22.2.3.3 PCR for Enterotoxin Gene of B. fragilis...........................................................................................314 22.3 Conclusions and Future Perspectives..............................................................................................................................314 References...................................................................................................................................................................................315

22.1  Introduction Gastrointestinal infections are the major cause of diarrhea, leading to the deaths of more than 2.2 million people per annum worldwide (WHO). In developing nations it has been estimated that approximately 1.8 billion episodes of diarrhea occur per year and 3 million children die under the age of 5 years, and 80% of these deaths are in children less than 2 years of age. In Southeast Asia and Africa, diarrhea is responsible for as much as 8.5% and 7.7% of all deaths respectively.1 The ninth and tenth revisions of the International Classification of Diseases (ICD-9 and ICD-10) classify diarrhea according to etiological agent, rather than symptom complex. Because a number of agents may cause either watery or bloody diarrhea, and because persistent diarrhea is defined by duration of illness rather than etiology, the ICD codes do not correspond neatly to the symptom complexes.2 Ninety percent of the acute diarrhea cases are of infectious origin and the remaining 10% or so are due to medications, toxic ingestions, ischemia, and other conditions. In developing nations, diarrhea is universally infectious in origin. Most infectious diarrheas are acquired by the feco-oral route. The infectious agents most often associated with diarrhea in young children in developing countries are as shown in Table 22.1.3 The serious health threats posed by waterborne pathogens, fecal contamination is one of the main factors in drinking water, in aquaculture and in recreational water. Traditionally, the evaluation of the health risk for waters contaminated by feces is obtained through the quantification of certain

microorganisms, and only rarely by the direct measurement of the real hazard, which is the actual concentration of the pathogens. These organisms are not necessarily ­source-specific; they can be hosted indistinctively by humans, farm animals or wildlife. Consequently, the typical organisms such as Salmonella enterica serovar typhi, Shigella spp., Hepatitis A virus, and Norwalk-group viruses, various serotypes of Salmonella, Escherichia coli, and Cryptosporidium spp. are the main part of fecal pollution In addition, B. fragilis bacteriophage and the F+ RNA coliphage further characterize and identified as a new tracing sources of fecal pollution.4 Bacteroides. fragilis is another emerging bacterial pathogen associated with diarrhea. Some strains of this species has been identified as a new virulence factor—referred to as enterotoxigenic B. fragilis (ETBF)—from fecal specimens of animals suffering from diarrhea in the United States by Myers et al.5 It has been suggested that ETBF strains can survive in municipal sewage and can be one of the causal agents of diarrhea. ETBF strains were later isolated in Poland, Italy, Japan, and Bangladesh, from intestinal and extraintestinal sources.6–9 In this chapter a molecular detection method for B. fragilis as well as ETBF and its biology, epidemiology, and pathogenesis will be discussed.

22.1.1  Biological Characteristics B. fragilis is an anaerobic small Gram-negative bacillus and coccobacillus in the genus Bacteroides, and is among the important components of the normal human flora, in the oral cavity, gastrointestinal tract, and the vagina.10 Its classification 307

308

Molecular Detection of Foodborne Pathogens

Table 22.1 Pathogens Frequently Identified in Children with Acute Diarrhea in Treatment Centres in Developing Countries Pathogen Viruses Bacteria

Protozoans No pathogen found

% of Cases Rotavirus Enterotoxigenic E. coli Shigella Campylobacter jejuni Vibrio cholerae O1 Salmonella (non Typhoid) Enteropathogenic E. coli Cryptosporidium

15–25 10–20 5–15 10–15 5–10 1–5 1–5 5–15 20–30

had undergone many changes since Veillon and Zuber named their isolates B. fragilis, and B. fusiformis. Castellani and Chalmers changed it to genus Bacteroides in 1994. After a lot of debate over the years, the first general agreement on taxonomy and classification was reached at the meeting of International Commission for Systematic Bacteriology SubCommittee for Gram-negative bacillus at Lille in 1967, when new principles of classifications were defined.11 Physiological analysis of this genus revealed considerable heterogeneity with regard to their biochemical properties, indicating these bacteria did not represent a true phylogenetic grouping. With the advent of phylogenetic analysis techniques, several investigators have tried to redefine this group of bacteria using physiological characteristics serotyping, bacteriophage typing, lipid analysis, oligonucleotide cataloging, and 5S–16S rRNA sequence comparisons.12–20 Based on this information, the original Bacteroides members have been partitioned into three genera: Bacteroides, Prevotella, and Porphyromonas.21–23 The Bacteroides are found predominantly in the colon of mammals, while the Prevotella and Porphyromonads generally are associated with the oral cavity and the rumen. In Bergeys Manual the Bacteroidaceae family is divided into three genera: Bacteroides, Fusobacterium, and Leptotricha. The genus Bacteroides consists of 22 species in five groups: (1) B. fragilis included most of the species described by Eggerth and Gagnon and was divided into five subspecies. (2) Phenotypically similar strains that were inhibited by bile included B. rumnicola, B. oralis, B. ochraceus and B. amylophilus. (3) A group of six species that did not produce succinic acid, but otherwise unrelated. (4) Nonsaccharolytic nonpigmented strains. (5) B. melaninogenicus produced black-pigmented colonies on laked blood agar and was further subdivided into three subspecies.11 Presently Bacteroides is characterized as follows: (i) obligately anaerobic, Gram-negative; (ii) saccharolytic, producing acetate and succinate as the major metabolic end

products; (iii) contain enzymes of the hexose monophosphate shunt-pentose phosphate pathway; (iv) have a DNA-base composition in the range 40–48 mol% GC; (v) membranes contain sphingolipids, and contain a mixture of long-chain fatty acids, mainly straight chain saturated, anteiso-methyl, and iso-methyl branched acids; (vi) possess menaquiones with MK-10 and MK-11 as the major components; and (vii) contain meso-diaminopimelic acid in their peptidoglycan.24 On gas liquid chromatography (GLC), it shows a peak at propionic acid in contrast to Fusobacterium, which shows a peak at butyric acid. They are moderately pleomorphic, but long filamentous, bizarre shapes, L-forms, and spheroplasts are rare.11 It is one of the most commonly isolated anaerobic organisms from infected tissues. Bacteroides are of primary importance to humans when they cause opportunistic and extraintestinal infections. They can cause a wide spectrum of clinical manifestations including periodontal disease, postaspiration pleuropulmonary infection, genital tract infections in females, and intraabdominal abscess.10 A subgroup of this species which may be important as an etiological agent of acute diarrheal illness in humans has now been identified.25 The role of Bacteroides as an etiological agent for diarrhea was not considered until 1984, when Meyer and his coworkers first reported that some strains of B. fragilis were associated with acute disease in newborn lambs by producing a factor with enterotoxigenic activity.5 Three years down the line evidence started to suggest that ETBF may have a role in human diarrhea when in close contact with animals. These studies targeted the children population of developing countries.26 Later, two studies were conducted in the developed nations: significant association of ETBF with childhood diarrheal disease was found in the United States but in Italy, though the presence of ETBF was seen, it has not reached statistical significance.9,25–27 Later on, many studies supported the fact that ETBF is a causative agent of self-limiting diarrhea. Recent studies have shown that the rate of ETBF carriage is high in both adults and children, regardless of whether diarrhea is absent or present. It has been suggested that ETBF may be endemic in communities.28

22.1.2 Epidemiology B. fragilis constitutes about 1–2% of normal colonic microflora and although rare, it is a significant causative agent in childhood diarrhea. In case control studies, ­enterotoxin-producing strains of B. fragilis have been implicated as a cause of watery diarrhea limited to children 1–5-years-old. In this age group it is responsible for 5–20% of diarrhea cases.10 It has also been shown that 6.5% of healthy persons could harbor ETBF without any clinical illness. Many prospective and epidemiological studies were conducted in different countries to understand the pathogenic role of ETBF and its association with diarrheal disease by using cytotoxicity and PCR assay. Sack et al.9,25 reported 12% isolation rate of ETBF strain in Native Americans and 9% in Bangladeshi children, compared with 6% of controls. Joaquin et al. found a strong association between diarrheal disease and ETBF in children with

Bacteroides

the isolation rate of 4.8%, whereas B. fragilis were recovered in 32.1% of children with diarrhea in an urban setting in the United States. Pantosti et al.7 reported 17% isolation rate and found that in Italy the rate of ETBF carriage is high in both adults and children and B. fragilis was found in 43% of children.7 In another study, ETBF was isolated from 13.2% of patients with inflammatory bowel disease.29 In age-matched controlled studies performed in Japan, ETBF was isolated from 14.9% of 114 children aged 1–14 years with antibioticunassociated diarrhea (AUD).30

22.1.3  Pathogenesis B. fragilis forms less than 10% of the Bacteroides in normal human feces, yet it is by far the most common species of Bacteroides to be isolated from infections related to intestine. It thus appears to be particularly pathogenic to human beings. This has been attributed to the presence of a polysaccharide capsule or to the action of one or more of the extracellular or membrane-associated enzymes that it forms: proteinases, including collagenase, fibrinolysin, hemolysin, neuraminidase, phosphatase, DNAase, hyaluronidase, chondroitin sulphatase, and heparinase. Exotoxins that are so important in disease caused by Clostridium spp. do not contribute significantly to the pathogenesis of infection by anaerobic rods. An exception to this is an enterotoxin-producing B. fragilis that has been associated with diarrhea.10 Recent evidence suggest its role in antibiotic-associated diarrhea (AAD). In age-matched controlled studies performed in Japan, ETBF was isolated from 6.5% of 108 children aged 1–6 years with AAD.30 In a study conducted by Pituch et al.,31 ETBF strains were cultured from 6.7% of investigated fecal samples. In a very recent study, it was shown that out of 50 stool samples of AAD, nine showed the presence of ETBF.31 Therefore, it shows that the role of ETBF in various clinical disorders predisposing to diarrhea cannot be underrated.

22.1.4 Enterotoxin: Virulence Factor of B. fragilis The toxin is the major virulence factor in the causation of diarrheal disease. The enterotoxin of B. fragilis is an extracellular, heat labile metalloprotease with a molecular weight of about 20-kDa that have both cytotoxic and secretory components. Metalloproteases have emerged as an important virulence factor in a number of diverse pathogenic organisms including bacteria and fungi. It has been implicated in a variety of virulence functions, including adhesion, tissue invasion, activation of protoxins and other proenzymes and cytotoxicity. Pseudomonas aeruginosa and Aspergillus fumigatus produce elastases that contribute to their invasiveness. Vibrio cholerae produces a hemagglutinating metalloprotease that nicks and activates cholera toxin. Staphylococcus aureus produces it for activation of V8 protease. Streptococcus sanguis and Streptococcous pneumoniae produce an extracellular metalloprotease that hydrolyze IgA and helps in adherence. Recently Clostridium botulinum and Clostridium tetani have

309

also shown the presence of Zn-dependent metalloprotcase. The ubiquity of metalloproteases in nature including many endogenously produced by the colonic microflora gives sufficient evidence about acquisition of enterotoxin by the B. fragilis.31–36 The enterotoxin of B. fragilis is an also important virulence factor inside and outside the intestinal lumen. It produces extensive tissue damage in the intestinal mucosa in vivo and increases bacterial internalization by enterocytes. The enterotoxin is tight junction specific; causes rounding, swelling, and pyknosis of enterocytes in culture; and induces a fluid response in ligated intestinal loops and a cytotoxic response in the HT-29 colon cell line. This toxin binds to a specific intestinal epithelial cell receptor and stimulates cell proliferation, which is dependent, in part, on E-cadherin degradation and β-catenin–T-cell-factor nuclear signaling. γ-Secretase is an intramembrane cleaving protease and is a positive regulator of E-cadherin cleavage and a negative regulator of β-catenin signaling. Toxin induces step-wise cleavage of E-cadherin, which is dependent on metalloprotease and γ-secretase toxins. This proteolytic event is hypothesized to trigger the reported F-actin rearrangement and pathophysiologic sequelae stimulated by the enterotoxin.37 Since the recognition of this enteric pathogen, comparison of the biologic activities of crude culture supernatants of ETBF strains in LLIL and the HT29/C1 cell assay have revealed variable responses with some strains classified as high BFT (B. fragilis toxin) producers and others as moderate or low BFT producers. Three distinct nucleotide sequences for bft were first reported for ETBF strains VPI 13784 (lamb), 86-5443-2-2 (piglet) and Korea 419 (human). Alignment of these sequences revealed 87–96% identity in the predicted protein sequences of the mature BFT proteins, termed BFT-1 (VPI 12784), BFT-2 (86-5443-2-2), and BFT-3 (Korea 419). All BFTs appear to be structurally similar.38 BFT is synthesized as a 44-kDa precursor (397 amino acid residues) containing the following three consecutive peptide domains: (i) a presignal sequence (18 amino acid residues), (ii) a propeptide (193 amino acid residues), and (iii) a mature protein (186 amino acid residues). The 44-kDa precursor protein is processed to a 20-kDa mature BFT that is secreted into the culture supernatant. The 20-kDa mature BFT contains the zinc-binding metalloprotease motif (H348–H358) and a methionine residue seven amino acids C terminal to the zincbinding metalloprotease motif, typical of the matrix metalloprotease (MMP) family. This may explain to some extent that various biologic responses to culture supernatants of ETBF strains were differences in the protein toxin secreted by these strains.39 Alternatively, differences in the efficiency of synthesis and secretion of BFT by different strains and/ or accessory virulence genes present in some, but not all, ETBF strains may account for the strain-dependent biologic activities detected to date. Recent studies have demonstrated that BFT induces the expression of interleukin-8 (IL-8) in human intestinal epithelial cells (HT29, T84, and Caco-2). It has also been suggested that there could be significant association between detection of the bft gene in stool specimens

310

Molecular Detection of Foodborne Pathogens

7 kb region ORF 10

11 12

Int2

13

15 16

rteB rteA satG bexA ORFs oriT 20–24

CTn9343 tnpB tnpA1 Probes:

14

4

ORFs 25–29

Transfer ORFs 35–50

bfmC 5

tnpA2

12 kb region

6

7

8

9

10

11 12

Figure 22.1  Schematic map of CTn9343. (From Buckwold, S.L., Shoemaker, N.B., Sears, C.C., and Franco, A.A. Appl. Environ. Microbiol., 73, 53, 2007.)

of inflammatory bowel disease patients. In a recent study, a ­collection of ETBF and nontoxigenic B. fragilis (NTBF) strains, it was found that bft is located in a ~6-kb region present only in ETBF strains (Figure 22.1). Sequence analysis of this 6-kb region revealed that, in addition to bft, this region contains another metalloprotease distinct from bft. Together, these observations indicate that bft is contained in a pathogenicity island (PAI). Based on the presence of the PAI and its flanking region, three major populations of B. fragilis strains were identified: (i) pattern I strains, containing the PAI and its flanking region, which are all ETBF strains; (ii) pattern II strains, lacking the PAI and its flanking regions, which are all nontoxigenic B. fragilis (NTBF) strains; and (iii) pattern III strains, containing the flanking region but lacking the PAI, which are all NTBF strains. The G-C content of the B. fragilis PAI (35%) and of the flanking DNA (47–50%) differs greatly from that reported for the B. fragilis chromosome (43%) suggesting that the PAI and its flanking region are two distinct genetic elements originating from different organisms. Based on these results, it was hypothesized that ETBF strains may have evolved by horizontal transfer of these two genetic elements into a pattern II NTBF strain. It was recently determined that the genetic element flanking the PAI in ETBF 86-5443-2-2 (pattern I; contains the PAI) and a related genetic element in NTBF NCTC 9343 (pattern III; lacks the PAI) are putative conjugative transposons (CTns), designated CTn86 in strain 86-5443-2-2 and CTn9343 in strain NCTC 9343. In contrast to most of the CTns detected in Bacteroides spp., CTn86 and CTn9343 do not carry the tetracycline resistance gene tetQ, and the excision of the transposons from the chromosome is not regulated by tetracycline. Based on these characteristics, sequence homology, and the proposed mechanism of transposition, CTn86, and CTn9343 may be members of a new family of CTns in Bacteroides spp. not described previously.

22.1.5 Role of Molecular Methods for B. Fragilis The acceptance of B. fragilis as a new diarrheagenic agent in animals led directly to the question whether it is possible to isolate the ETBF strains from the humans gut flora and whether they could be the diarrheagenic agent.26 First observations gave a positive answer when strains were tested in lamb ileal loop (LIL) and on adult rabbit’s ceca. The bacterium colonized the intestinal tract with a predilection for

the large intestine, and diarrhea occurred. The disease was characterized by watery diarrhea and dehydration.40 Identification of enterotoxin production by culturing on selective medium (BBE) and by testing the isolates for presence of enterotoxin by cytotoxic assay with HT 29/C1 cells or LIL is cumbersome as a routine diagnostic laboratory method (due to animal ethical issues). Therefore scientists in this field considered molecular techniques for the development of DNA probes, characterization of enterotoxigenic genes or genus specific genes (in case of failure of isolation by culture methods). Initially to overcome this problem, DNA probes (e.g., pBFII-4, pBFII-5, and pBFII-6) have been developed with specificity for B. fragilis, which accounts for about half of all isolates from clinical specimens containing anaerobes. The limit of detection for these probes was 106 bacteria. The probes could detect B. fragilis in blood culture medium and in mixed cultures with other Gram-negative bacteria. Attempts to use biotin-labeled DNA probes instead of 32Plabeled probes were not successful because the Bacteroides spp. extracts contained material that bound the streptavidinperoxidase detection reagent. Later it was found to be less sensitive and specific. PCR allows the rapid and specific detection of a wide range of bacterial species and it has become a key procedure for detecting microorganisms. PCR targeting the 16S rRNA was developed. The series of probes were developed and consist of a universal bacterial probe, a Gram-positive probe, a Bacteroides-Flavobacterium probe, and two probes for other Gram-negative species. However, in terms of specificity it was not the test of choice. The direct detection of B. fragilis from clinical specimens is also examined using the PCR method for amplifying a specific fragment of the glutamine synthetase gene from B. fragilis.41 The test was found to be fairly helpful in the initial stages but detection was limited to a specific range of clinical specimens. The attention of the scientists was drawn towards another gene target present in B. fragilis. Neuraminidases are produced by many bacterial species including pathogens, e.g., Vibrio cholerae and Clostridium perfringens and indigenous bacteria such as a wide range of species within the family Bacteroidaceae. Neuraminidase enzyme is produced by all B. fragilis strains in vivo and in vitro at elevated levels. The neuraminidase gene nanH of B. fragilis has been partially sequenced. To date this is one of the methods of choice for detection of B. fragilis. We will discuss in detail

311

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the methodology adapted for the PCR of neuraminidase gene (genus specific) and enterotoxin gene for confirmation of ETBF strain from culture and directly from fecal specimen. The identification of this organism in clinical microbiology laboratory is sometime difficult, as it is a fastidious organism and takes 3–5 days when grown in culture. Moreover, B. fragilis are often unrecognized in fecal samples because of inappropriate transportation (within 2 h of collection), specific growth requirements and frequent coexistence with aerobic organisms, present in the fecal sample. Further processing for cytotoxic assay and rabbit ileal loop assay for enterotoxin test requires additional time. Due to ethical issues involved with animal experiments and nonavailability of rapid diagnostic technique, isolation of ETBF is generally overlooked. Therefore, it becomes essential to apply a rapid and sensitive molecular diagnostic technique to detect neuraminidase gene specific for B. fragilis and enterotoxin gene to detect ETBF directly from fecal samples of patients with diarrhea.

22.2  Methods 22.2.1 Reagents and Equipment General reagents • • • • • • • • • •

EDTA NaCl NaOH HCl Sodium dodecyl suphate (SDS) Tris base Boric acid Agarose Ethidium bromide (10 mg/ml) Loading dye

PCR reagents (see Table 22.2) • PCR buffer • dNTPs

• MgCl2 • Primers • DNA polymerase Equipment • • • • • • • • •

Thermal cycler Water bath pH meter Autoclave Gel electrophoresis tank with power pack Magnetic stirrer Pipettes Electronic balance Centrifuge

22.2.2  Sample Collection and Preparation Enterotoxigenic strains of B. fragilis are pathogenic, this bacterium is bile stimulated and extremely saccharolytic. Basically, choice of specimen depends on the site of infection. In cases of ETBF, a fecal specimen is the sample of choice. Fecal swabs are not considered because of excessive exposure to the deleterious effects of drying, the possibility of contamination during collection and the easy retention of microorganism within the fibers of the swab. The proper collection and transport of specimens for anaerobic culture cannot be overemphasized. Because indigenous anaerobes are often present in large numbers as normal flora on mucosal surfaces, even minimal contamination of a specimen can give misleading results. Collection and transportation of fecal specimen. Specimens must be protected from the deleterious effect of oxygen until they can be cultured. In proper anaerobic transport medium, anaerobic bacteria may be survive for up to several days. Fecal specimens should be collected in wide mouth containers and inoculated as soon as possible on respective media within 2 h of collection because it contains enzymes that degrade bacterial component.

Table 22.2 Commercially Available PCR Reagents for Preparation of Master Mix Final Concentration 1× 200 μM 1–3 mM

0.2–1.0 μM 2–2.5 U/1 μl 0.05–0 μg

Component

Purpose

PCR buffer

Keeps the master mix at an optimum pH favorable for PCR.

Deoxynucleotides (dNTPs-mix) MgCl2

Provides both energy and nucleosides for the synthesis of DNA. The master-mix must have equal concentrations of each nucleotide (dATP, dTTP, dCTP, dGTP) to avoid mismatches of bases. Mg++ ions are essential for DNA polymerase activity. Optimal conc. of Mg++ is required since suboptimal levels result in low yield of PCR products whereas excess of it may increase the yield of non-specific products and base-mismatches. Short pieces of DNA (20–30 bases) that bind to the DNA template allowing Taq DNA polymerase enzyme to initiate incorporation of the deoxynucleotides. Both specific and universal primers can be used. A heat stable enzyme that adds the deoxynucleotides to the DNA template.

Primers (forward and reverse) Ampli-Taq-DNA polymerase Template DNA Sterile water

The DNA which will be amplified by the PCR-reaction. Adds up to the final of reaction mix.

312

Specimen processing. Specimens can be processed on the open bench top with immediate incubation under anaerobic conditions. For B. fragilis fecal samples, streak on Bacteroides bile esculine agar (BBE), brain heart infusion agar (BHIA) supplemented with haemin and Vit.K(Diffco)and incubate anaerobically at 37oC for 24 h and 48–72 h. Anaerobiosis for the growth of B. fragilis and other anaerobes is generated by using ANOXOMAT System (MART Sint-Genesius-Rode, Belgium), at anaerobic gaseous atmosphere (80% N2, 10% H 2 and 10% CO2) in an anaerobic jar. BBE agar is useful for rapid isolation and presumptive identification of B. fragilis group. The medium contains 100 g/ml of gentamicin to inhibit most of aerobic organisms and 20% bile to inhibit other anaerobes. As differential agent, esculin, and iron have been incorporated into the agar to aid in detecting the esculin-positive, B. fragilis group organisms. BBE agar is also a useful medium due to H2S production. Other organisms that may grow on this medium are (Fusobacterium mortiferum, F. varium, gentamycin resistant Entreobacteriaceae, enterococci, pseudomonads, staphylococci, and yeast. However, the colony size of the facultative anaerobic organism is usually less than 1 mm in diameter. Selective media (BBE) may be examined after 24 hrs of incubation, as selective organism grows rapidly. Examination of culture. After incubation, observe colonial characteristics and record detailed colony description, i.e., color, hemolysis, fluorescence, pigment, shape, and size with peripheral striations: crenate/entire edge. On BBE agar they hydrolyze esculin, blackening the agar. When all colony types are confirmed as pure anaerobes, do Gram-stains and further subculture on BHIA with special potency disc (kanamycin 1 mg, colistin 10 μg, vancomycin 5 μg, sodium polyanetho sulfonate 1 mg, penicillin 2 units). A zone size of >10 mm is considered as sensitive and <10 mm is considered as resistant. The special potency antibiotic disc pattern (colistin-resistant and vancomycin-sensitive) will demonstrate the Gram-positivity of isolates. Pure colonies of B. fragilis on BBE and BHIA show circular, low convex 1–3 mm diameter colonies with an entire edge. They are usually smooth, shiny, translucent, or semi-opaque. They are nonhemolytic. A granular precipitate around the colonies is an important characteristic of B. fragilis sp. They are further identified by the use of conventional biochemical tests or by using API 20A/API 32 A system (bioMeruix). Some of the species within the group are very similar biochemically and therefore difficult to differentiate. Biochemical tests. It is catalase negative and grows well in 20% bile and is resistant to kanamycin, vancomycin, and colistin by special potency identification antibiotic discs. They hydrolyze esculin and show fermentation of three major sugars namely, glucose, sucrose, and lactose. In fact the above-mentioned methods are quiet helpful, but due to the long processing time to complete the identification, these tests are no longer considered an option. Therefore, rapid tests based on molecular methods are the techniques of choice.42

Molecular Detection of Foodborne Pathogens

Determination of enterotoxin production. In 1984 Myers et al. tested enterotoxin production by ligated animal loops in isolated B. fragilis, which was difficult and complicated for the diagnostic laboratory.5 Weikel et al. described a culture supernatant method for enterotoxin determination in 1992.43 It was a cytotoxicity test on human colon carcinoma cell lines (HT 29/C1), that proved a very useful in vitro assay for Enterocytotonic activity. In this test ­sub-confluent cells treated with bacterium free culture supernatant of ETBF strains developed specific morphologic changes such as loss of cell-to-cell attachment rounding, swelling, and in some cases pyknosis. The morphological changes has been initially visible after 1 h of treatment and progressed over at least the first 24 h. To avoid taking culture over a long time and further cytopathic activity, some basic molecular methods have been introduced. These methods shorten the procedure by detecting the DNA directly from clinical specimens or from B. fragilis isolates (Figure 22.2).

22.2.3  Detection Procedures Scientists have been searching for more rapid and efficient methods to detect and characterize microorganisms. Recently introduced into food and clinical microbiology, molecular techniques provide some of the most powerful tools. These methods have revolutionized the microbiology laboratory and have changed the process of detection, characterization, and quantification of microorganisms directly from clinical specimens and from cultures.43 Development of molecular assays is particularly useful for detection of fastidious or uncultivable microorganisms and to determine the cause of significant outbreaks. Nowadays, these methods are more reliable, more accurate, easier to apply and, once established are less time-consuming. Molecular methods in microbiology can be divided in three categories: (i) detection of microorganisms without using nucleic acid amplification such as in situ nucleic acid detection; (ii) target detection by restriction fragment length polymorphism (RFLP) analysis and pulse-field gel electrophoresis (PFGE) by purification of nucleic acids. These methods have been used extensively to determine the relatedness among microorganisms (i.e., strain typing), which have become a critical tool for the hospital and public health epidemiologist; (iii) detection, characterization and, in some instances, quantification of microorganisms by using a variety of nucleic acid amplification methods (PCR, LCR, etc.) for in vitro amplification and detection of a specific nucleic acid sequence of a gene. Below we present procedures for extraction of DNA ETBF in culture and directly from fecal samples and their subsequent identification by molecular techniques. 22.2.3.1 DNA Extraction Phenol-chloroform extraction. Suspend one or two pure colonies from culture plate in 100 µl of Tris-EDTA [10 mM Tris-HCl pH 7.5, 1 mM EDTA], and 200 µl of lysis buffer [200 mM NaCl, 20 mM EDTA, 20 mM Tris-HCl pH 8.0,

313

Bacteroides

4% (w/v) SDS, 1 mg/ml proteinase K] in a microcentrifuge tube. Incubate in a water bath at 60oC for 2 h. After cell lyses, extract DNA by the phenol-chloroform method (25:24:1) with the following step-wise protocol:

Keep at –20oC overnight Centrifuge at high speed for 15 min at room temperature

Take 100 µl of clinical sample (stool/pus) in a 1.5-ml mi crocentrifuge tube

Decant supernatants and wash with 70% alcohol and spin for 15 min

Add equal volume of TE buffer (100 µl) [10 mM Tris-HCl pH7.5, 1 mM EDTA] Mix with 200 µl of lysis buffer [200 mM NaCl, 20 mM EDTA, 20 mM Tris-HCl pH 8.0, 4% (w/v) SDS, 1mg/ml proteinase K]

Decant supernatant and dry the pellet at 37oC for 1 h Dissolve DNA in 30 µl TE buffer and store at 4oC or use immediately for PCR

Keep in a water bath (60°C) for 1 h Add phenol-chloroform (equal volume) and spin at 6,000 rpm for 10 min (repeat this step twice) Transfer supernatant to a fresh tube and add cold absolute alcohol (two volumes) and 4 M NaCl (1/10th of the volume), invert the tubes a few times

Boiling method. Extraction by boiling is a simple and rapid method of DNA-preparation for PCR. Briefly, colonies of isolated and identified B. fragilis are picked up with sterile loop and suspended in sterile phosphate buffer saline (PBS) pH 7.2 in a 1.5-ml microcentrifuge tube, washed with 1× PBS and centrifuged at 4,000×g for 1 min (two to three times). The pellet is resuspended in 50 μl sterile double distilled water and boiled for 3–5 min. After a short spin (30 sec) in a centrifuge (eppendorf), the supernatant (40 μl) containing DNA is collected in new tube.

Clinical specimen RCM

Bacteroides Bile Esculin

BHIA

Incubate 24 h solution

Agar (BBE)

(supl.Vit K and Heamin Solution) (Incubate 24–48 h)

Gram stain Aerobic (MC)

Anaerobic

Aerobic

sub inoculate

(BBE/BHIA)

(MC)

Anaerobic (BBE/BHIA)

(MC, BA) G+

G– G+

(No Anaerobe)

G–

G+

(No Anaerobe)

G–

Gram stain

(No Anaerobe) KVCPS

Biochemical test API 20A

Pure colony isolates

Gram Special potency disc stain (KVCPS)

Biochemical test API 20A

Figure 22.2  Flow chart for processing anaerobes. RCM, Robertson cooked meat medium; BHIA, brain heart infusion agar; MC, MacConkey agar; BA, blood agar.

314

Molecular Detection of Foodborne Pathogens

Culture the strain on BHIA Agar/Robertson Cooked Meat medium (RCM) Take a few colonies and suspend in sterile PBS (1 ml) or 1 ml of cultured RCM and spin at 4,000×g for 1 min Discard the supernatant and add PBS for washing the cells (repeat this step two or three times) Resuspend the pellet in 50 µl sterilized distilled water and boil at 100oC for 3 min Short spin (30 sec), collect the supernatant (containing DNA) in another microcentrifuge tube for PCR. Extraction of DNA from fecal samples. Homogenize one loop full of a fresh stool sample (within 10 min of collection) by adding glass beads (two to three autoclaved) and 200 μl sterile PBS or normal saline and vortex for 1 min. Keep this tube in racks for 1 min and after larger fecal matter has settled down, take 100 µl of supernatant from that sample in a 1.5-ml tube. Add 100 µl of TE buffer [10 mM Tris-HCl pH 7.5; 1 mM EDTA] and 200 µl of lysis buffer [200 mM NaCl, 20 mM EDTA, 20 mM Tris-HCl pH 8.0, 4% (w/v) SDS, 1 mg/ml proteinase K]. Incubate in a water bath at 60oC for 2 h. After cell lyses, extract DNA by the phenolchloroform method (25:24:1) and then proceed for PCR. Take one loop full of a fresh stool* sample (i.e., within 10 min of collection) in a microcentrifuge tube Add 200 μl sterile PBS or NS and homogenize by adding glass beads (two to three autoclaved) and vortex for 1 min Keep the tubes in racks for 1 min and after the fecal matter settles, take 100 µl of supernatant in a 1.5-ml microcentrifuge tube Add 100 µl of TE buffer [10 mM Tris-HCl pH 7.5; 1 mM EDTA] and 200 µl of lysis buffer [200 mM NaCl, 20 mM EDTA, 20 mM Tris-HCl pH 8.0, 4% (w/v) SDS, 1 mg/ml proteinase K] Incubate in a water bath at 60 C for 2 h o

After cell lyses, extract DNA by the phenol-chloroform method (25:24:1) and then proceed with PCR (Extraction should be started with spiked stool sample). 22.2.3.2  PCR for Neuraminidase Gene Neuraminidase enzyme is produced by all B. fragilis strains at elevated levels in comparison to other * If it is a diarrheal sample then do not add saline, use 100 µl of sample for DNA extraction.

Bacteroides sp. Considered as genus specific the PCR assay for detection of neuraminidase gene of B. fragilis is both a suitable and alternative option when failure of cultivation of this bacterium from fecal specimens occurs. It is carried out by using primers (forward GAI 11: 5′GCCGGTCAGAATGGGAGTAGGAGACC-3′; and reverse GAI 12: 5′-CCCGACGAGCCGGACCTTGCAACAGA-3′), designed to amplify a 262-bp sequence, specific for neuraminidase gene as reported by Jatwani et al.8 The PCR reaction mixture consists of 50 pM of each primer, 100 μM each dNTPs, 1 U of Taq DNA polymerase, 1× PCR buffer containing 1.5 mM MgCl2 and 5 μl of DNA template in 25 μl of reaction volume. PCR cycle consists of one cycle of 94oC for 1 min; 35 cycles of 94oC for 1 min, 62oC for 1 min, and 72oC for 1 min; one cycle of 72oC for 5 min. The amplified PCR product (262 bp) is detected under UV, after staining with ethidium bromide. After the establishment of ETBF as a causative organism in diarrhea, it has now been well understood that mere detection of B. fragilis in the stool specimen is not sufficient. Therefore, currently PCR-based techniques are targeting the enterotoxin gene for confirmation of the presence of ETBF. 22.2.3.3  PCR for Enterotoxin Gene of B. fragilis DNA extraction from clinical specimens is similar to the process used for detection of neuraminidase gene. PCR assay for detection of enterotoxin gene of B. fragilis is carried out by using primers (forward BF1: 5′-GACGGTGTATGTGAT TTGTCTGAGAGA-3′ and reverse BF2: 5′-ATCCCTAAGA TTTATTATCCCAAGTA-3′), designed to amplify a 294-bp sequence, specific for neuraminidase gene as reported by Pantosti et al.7 The reaction mixture contained dNTPs 100 μM each, 50 pM of each primer, 1 U of DNA polymerase and 10 μl of DNA template in a final volume of 25 μl of 1× PCR buffer containing 1.5 mM MgCl2. Samples are subjected to one cycle of 94oC for 3 min; 35 cycles of 94oC for 1 min, 58oC for 1 min, and 72oC for 1 min; one cycle of 72oC for 5 min. The PCR product is subjected to electrophoresis in 1.6% agarose gel, stained with ethidium bromide and examined under Gel Doc System.

22.3 Conclusions and Future Perspectives Diarrhea is major cause of mortality in infants and children in Asia and Africa. According to WHO, diarrhea leads to the deaths of more than 2.2 million people per annum worldwide. It is mainly caused by bacteria both aerobic and anaerobic, viruses, and parasites. Among bacteria, most of the studies are focused on the isolation of enterotoxigenic E. coli, C. difficle, Helicobacter spp., Shigella spp. etc., but there are groups of organism especially B. fragilis that have been overlooked due to lack of awareness and suboptimal methods of diagnosis. Therefore, this chapter highlights the role of molecular methods especially PCR in detection of enterotoxigenic

315

Bacteroides

strains of B. fragilis. On the basis of culture technique it is difficult to differentiate enterotoxigenic strains of B. fragilis from nonenterotoxigenic strains of Bacteroides. Molecular methods will be very useful not only in diagnosis but also in epidemiologic studies of Bacteriodes spp. Interested readers are advised to refer to specific manuals like Sambrook et al.46 and obtain relevant literature from manufacturers. In general this chapter contains well documented and invaluable information which will help establish PCR techniques for B. fragilis as well as ETBF. A number of upcoming technologies show potential for further improving the laboratory diagnosis of B. fragilis and other microbial pathogens. For nucleic acid amplification, reverse transcription-PCR, broad range PCR, multiplex PCR, nested PCR, and real-time PCR can be exploited. For post amplification analysis, gel electrophoresis, Southern blot analysis, enzymatic detection of amplified products, reverse hybridization, DNA sequencing, sequences by synthesis (pyrosequencing) and microarray analysis can be utilized. For strain typing, pulsed-field gel electrophoresis, PCRRFLP and REP-PCR can be applied. The number of molecular applications for the detection and characterization of all types of microorganism increases each year, with many commercial venders seeking FDA approval for their application. These amplification include numerous signal-amplification methods, which in many instances may be not as sensitive as PCR, but are simple to use, may be less expensive and are not burdened with the specter of potential amplicon contamination. Nucleic acid amplification assays, and the methods to detect the product of amplification have evolved rapidly in the past decade, culminating in the exiting application of real-time PCR and similar technologies. Finally, as it is difficult to analyze each and every sample for various pathogens using standard molecular techniques, we need a all inclusive approach by developing and utilizing DNA microarray chips which will be helpful in the detection of numerous pathogens simultaneously without the need of sophisticated equipment. These diarrheagenic agents are also a very important component of bioterrorism, as water is the transmission vehicle. In such a scenario the role of chipbased technology becomes more important. They are sensitive, specific, rapid, easy to use and above all they can be a very useful tool in the field.



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1. Mudur, G. India’s burden of waterborne diseases is underestimated. BMJ, 326, 1284, 2003. 2. Cary, B. Diarrhoeal diseases. In Christopher J.L. Murray (ed.), Global Epidemiology of Infectious Diseases, pp. 1–28. World Health Organization, Geneva, 2004. 3. Park, K. Acute diarrhoeal diseases: II. Intestinal infections. Epidemiology of communicable diseases. In Textbook of Preventive and Social Medicine, 17th Edn, India, 2002 (reprinted in in 2003). 4. Scott, T.M. et al. Microbial source tracking: current methodology and future directions. Appl. Environ. Microbiol., 68, 5796, 2002.

















5. Myers, L.L. et al. Bacteroides fragilis: a possible cause of acute diarrheal disease in newborn lambs. Infect. Immun., 44, 241, 1984. 6. Meisel-Mikołajczyk, F., Pituch, H. and Rouyan, G.S. Detection of enterotoxigenic Bacteroides fragilis (ETBF), among strains isolated between 1976 and 1995 in Poland. Acta Microbiol. Pol., 45, 187, 1996. 7. Pantosti, A. et al. Detection of enterotoxigenic Bacteroides fragilis and its toxin in stool samples from adults and children in Italy. Clin. Infect. Dis., 24, 12, 1997. 8. Jotwani, R. et al. Detection of Bacteroides fragilis in clinical specimens by polymerrase chain reaction amplification of the neuraminidase gene. Curr. Microbiol., 31, 235, 1995. 9. Sack, R.B. et al. Isolation of enterotoxigenic Bacteroides fragilis from Bangladeshi children with diarrhea: a controlled study. J. Clin. Microbiol., 32, 960, 1994. 10. Tzianabos, A.O. and Kasper, D.L. Anaerobic Infections: General Concepts. Principles and Practice of Infectious Diseases, 6th Edn, Vol. 2. Gerald Mandell, MD, University of Virginia, Health Sciences Center, Charlottesville. 11. Shah, H.N., Gharbia, S.E. and Olsen, I. Bacteroides, Prevotella, and Porhyromonas. In S. Peter Borriello, Patrick R. Murray and Guido Funke MD, Bacteriology, 10th Edn, Vol. 2, pp.1913–1938. Hodder Arnold, London. 12. Holdeman, L.V.R., Kelly, R.W. and Moore, W.E.C. Genus I. Bacteroides Castellani and Chalmers 1919. In Krieg, N.R. and Holt, J.G. (eds.), Bergey’s Manual of Systematic Bacteriology, Vol. 1. The Williams and Wilkins Co., Baltimore, MD, 1999. 13. Lambe, D.W. Jr. Determination of Bacteroides melaninogenicus serogroups by fluorescent antibody staining. Appl. Microbiol., 28, 561, 1974. 14. Booth, S.J. et al. Bacteriophages of Bacteroides. Rev. Infect. Dis., 1, 325, 1979. 15. Miyagawa, E., Azuma, R. and Suto, T. Distribution of sphingolipids in Bacteroides species. J. Gen. Appl. Microbiol., 25, 41, 1979. 16. Paster, B.J. et al. A phylogenetic grouping of the Bacteroides, cytophagas, and certain flavobacteria. Syst. Appl. Microbiol., 6, 34, 1985. 17. Paster, B.J. et al. Phylogeny of Bacteroides, Prevotella, and Porphyromonas spp. and related bacteria. J. Bacteriol., 176, 725, 1994. 18. Johnson, J.L. Taxonomy of the Bacteroides. I. Deoxyribonucleic acid homologies among Bacteroides fragilis and other saccharolytic Bacteroides species. Int. J. Syst. Bacteriol., 28, 245, 1978. 19. Van den Eynde, H. et al. 5S ribosomal ribonucleic acid sequences in Bacteroides and fusobacteria: Evolutionary relationships within these genera and among Eubacteria in general. Int. J. Syst. Bacteriol., 39, 78, 1989. 20. Weisburg, W.G. et al. Natural relationship between Bacteroides and Flavobacteria. J. Bacteriol., 164, 230, 1985. 21. Shah, H.N. and Collins, M.D. Proposal to restrict the genus Bacteroides (Castellani and Chalmers) to Bacteroides fragilis and closely related species. Int. J. Syst. Bacteriol., 39, 85, 1989. 22. Shah, H.N. and Collins, M.D. Prevotella, a new genus to include Bacteroides melaninogenicus and related species formerly classified in the genus Bacteroides. Int. J. Syst. Bacteriol., 40, 205, 1990. 23. Shah, H.N., and Collins, M.D. Proposal for reclassification of Bacteroides asaccharolyticus, Bacteroides gingivalis, and Bacteroides endodontalis in a new genus. Porphyromonas, Int. J. Syst. Bacteriol., 38, 128, 1988.

316 24. Shah, H.N. The genus Bacteroides and related taxa. In Balows, A. et al. (eds), The Prokaryotes, 2nd Edn, Vol. 4, pp. 3593–3607. Springer-Verlag, New York, 1992 25. Sack, R.B. et al. Enterotoxigenic Bacteroides fragilis; epidemiologic studies of its role as a human diarrhoeal pathogen. J. Diarr. Dis. Res., 10, 4, 1992. 26. Myers, L.L. et al. Isolation of enterotoxigenic Bacteroides fragilis from humans with diarrhea. J. Clin. Microbiol., 25, 2330, 1987. 27. Joaquin, S. et al. Association of Bacteroides fragilis with childhood diarrhea. Scand. J. Infect. Dis., 27, 211, 1994. 28. Sears, C.L. et al. Enterotoxigenic Bacteroides fragilis. Clin. Infect. Dis., 20 (Suppl 2), S142, 1995. 29. Sharma, N. and Chaudhry, R. Rapid detection of enterotoxigenic Bacteroides fragilis in diarrhoeal faecal samples. Indian J. Med. Res., 124, 575, 2006. 30. Kato, N. et al. Prevalence of enterotoxigenic Bacteroides fragilis in children with diarrhea in Japan. J. Clin. Microbiol., 37, 801, 1999. 31. Pituch, H. et al. Prevalence of enterotoxigenic Bacteroides fragilis strains (ETBF) in the gut of children with clinical diagnosis of antibiotic associated diarrhoea (AAD). Med. Dosw. Mikrobiol., 54, 357, 2002. 32. DasGupta, B.R., and Tepp, W., Protease activity of botulinum neurotoxin type E and its light chain: cleavage of actin. Biochem. Biophys. Res. Commun., 190, 470, 1993. 33. Galloway, D.R. Pseudomonas aeruginosa elastase and elastolysis revisited: recent developments. Mol. Microbiol., 5, 2315, 1991. 34. Häse, C.C., and Finkelstein, R.A. Bacterial extracellular zinccontaining metalloproteases. Microbiol. Rev., 57, 823, 1993. 35. Markaryan, A. et al. Purification and characterization of an elastinolytic metalloprotease from Aspergillus fumigatus and immunoelectron microscopic evidence of secretion of this enzyme by the fungus invading the murine lung. Infect. Immun., 62, 2149, 1994. 36. Schiavo, G. et al. Tetanus and botulinum-B neurotoxins block neurotransmitter release by proteolytic cleavage of synaptobrevin. Nature, 359, 832, 1992.

Molecular Detection of Foodborne Pathogens 37. Wu, S. et al. Bacteroides fragilis toxin stimulates intestinal epithelial cell shedding and γ-secretase-dependent E-cadherin cleavage. J. Cell Sci., 120, 3713, 2007. 38. Wu, S. et al. Diversity of the metalloprotease toxin produced by enterotoxigenic Bacteroides fragilis. Infect. Immun., 70, 2463, 2002. 39. Sears, C.L. et al. The C-terminal region of Bacteroides fragilis toxin is essential to its biological activity. Infect. Immun., 74, 5595, 2006. 40. Meisel-Mikołajczyk, F. et al. Antigenic properties of LPS extracted from Bacteroides fragilis enterotoxin producing strains. Acta Microbiol. Pol., 48, 153, 1999. 41. Kuritza, A.P. et al. DNA probes for dentification ofclinically important Bacteroides species. J. Clin. Microbiol., 23, 343, 1986. 42. Somer, J.H. et al. Advanced identification methods. In Wadsworth, K.T.L. (ed.), Anaerobic Bacteriology Manual, 6th Edn, pp. 81–99. 2002. 43. Weikel, C.S. et al. Human colonic epithelial cells, HT29/ C1, treated with crude Bacteroides fragilis enterotoxin dramatically alter their morphology. Infect Immun., 60, 321. Star Publishing Company, Balmonte, California,1992. 44. Hans, R.J. et al. Nucleic acid technique in diagnostic microbiology, Section A. In Collee, J.G. et al. (eds), A Practical Medial Microbiology of Mackie and McCartney, 16th Edn, pp. 205–242. Churchill Livingston, New york, 1991. 45. Winn, W. et al. Molecular microbiology. In Color Atlas and Textbook of Diagnostic Microbiology, 6th Edn, pp. 133–149. Lippincott Williams and Wilkins, Baltimore and Philadelphia, 2006. 46. Sambrook, J., Fritsch, E.F, and Maniatis, T. Molecular Cloning: A Laboratory Manual, 2nd Edn. Cold Harbor Laboratory Press, Cold Spring Harbor, NY, 1989. 47. Buckwold, S.L., Shoemaker, N.B., Sears, C.C., and Franco, A.A. Identification and characterization of conjugative transposons CTn 86 and CTn 9343 in bacteroids fragilis strains. Appl. Environ. Microbiol., 73, 53, 2007.

23 Brucella

Sascha Al Dahouk

RWTH Aachen University

Karsten Nöckler

Federal Institute for Risk Assessment

Herbert Tomaso

Friedrich Loeffler Institute

Contents 23.1 Introduction.....................................................................................................................................................................317 23.1.1 Taxonomy...........................................................................................................................................................317 23.1.2 Pathogenicity and Biology.................................................................................................................................318 23.1.3 Brucellosis in Animals and Men.......................................................................................................................318 23.1.4 Epidemiology of Brucellosis..............................................................................................................................319 23.1.5 Microbiological and Serological Methods in the Diagnosis of Brucellosis .................................................... 320 23.1.5.1 Isolation and Identification.............................................................................................................. 320 23.1.5.2 Serological Tests.............................................................................................................................. 320 23.1.6 Molecular Diagnostic Techniques ................................................................................................................... 321 23.1.6.1 PCR.................................................................................................................................................. 321 23.1.6.2 Molecular Typing Methods.............................................................................................................. 322 23.2 Methods.......................................................................................................................................................................... 323 23.2.1 Sample Collection and Preparation.................................................................................................................. 323 23.2.2 Detection Procedures........................................................................................................................................ 323 23.2.2.1 Multiplex PCR................................................................................................................................. 323 23.2.2.2 Real-Time PCR................................................................................................................................ 324 23.3 Conclusions and Future Perspectives............................................................................................................................. 325 References.................................................................................................................................................................................. 327

23.1  Introduction Brucellosis is a (re-)emerging disease in animals and man and the most common bacterial zoonosis worldwide.1,2 Although many countries successfully established programs for eradication and control, about 500,000 human cases are annually reported mainly from endemic regions including the Mediterranean Basin, the Arabic Peninsula, the Middle East, North Africa, Asia, Central, and South America. The disease causing pathogen Brucella is transmitted from its animal reservoir to humans by direct contact with infected animals or more often through the consumption of raw food products of animal origin e.g., unpasteurized milk or cheese. After September 11th, 2001 further interest in Brucella aroused from its status as a class B bioterrorist agent.3 Brucella spp. were among the first biological agents weaponized in the 1950s because of their high contagiosity and due to the enormous economic impact of human and animal

brucellosis.4 Effective human vaccines are not available and countermeasures therefore have to rely on early diagnosis, adequate antibiotic treatment and the control of livestock and products of animal origin which may pose a risk for consumers.

23.1.1  Taxonomy Brucellae are Gram-negative, nonmotile, nonspore-forming and facultative intracellular coccobacilli.5 The 16S rRNA gene sequence of the genus indicates that Brucella belongs to the family of Rhizobiaceae within the alpha-2 subdivision of the class Proteobacteria closely related to Bartonella/ Rochalimaea, Ochrobacterium, and Agrobacterium.6,7 Historically, species and biovar (bv) classification of brucellae was based on natural host preference and phenotypic characteristics, i.e., CO2 requirement, H2S production, urease activity, dye-sensitivity, oxidative metabolic profiles, lysis by Brucella-specific bacteriophages, and agglutination with 317

318

monospecific antisera.8,9 In accordance with these ­features the genus Brucella currently comprises the six classical species B. melitensis bv 1–3 (primarily isolated from sheep and goats), B. abortus bv 1–6 and 9 (from cattle and other Bovidae), B. suis bv 1–3 (from pigs), bv 4 (from reindeer) and bv 5 (from small ruminants), B. canis (from dogs), B. ovis (from sheep), B. neotomae (from desert wood rats),10 two novel species of marine origin, B. pinnipedialis (formerly B. pinnipediae isolated from seals) and B. ceti (formerly B. cetaceae isolated from dolphins and whales),11,12 and the recently described species B. microti isolated from the common vole Microtus arvalis.13 Due to their high degree of DNA homology exceeding 90% for all Brucella spp, a monospecific genus has been proposed in which Abortus, Canis, Neotomae, Ovis, and Suis should be regarded as biovars of B. melitensis.14 However, in 2003 the Subcommittee on the Taxonomy of Brucella agreed to return to pre-1986 taxonomy and reapproved the six classical Brucella species with their recognized biovars.15

23.1.2  Pathogenicity and Biology Although brucellae do not display classical virulence factors such as exotoxins, cytolysins, proteases, fimbriae, flagellae, capsules, plasmids or lysogenic phages, virulent strains are able to evade host defence mechanisms, penetrate host cells, alter intracellular trafficking, avoid killing in lysosomes and modulate the intracellular environment.16 Especially in the early stage of infection the nonendotoxic smooth lipopolysaccharide (sLPS) of Brucella helps to protect these bacteria from anti-microbial attacks of the host immune system.17 Brucella LPS inhibits complement and cationic peptide-mediated killing and prevents the synthesis of immune mediators. Furthermore, the entry of Brucella in macrophages is partially dependent on the O-polysaccharide chain of LPS.18 LPS also interferes with MHC II antigen presentation keeping infected host cells from activating Brucella antigen-specific CD4+ T cells.19 However, cellular immunity is absolutely required for the control and resolution of Brucella infections in the host.20 Another key-factor involved in the modulation of cell binding and penetration is the two-component regulatory system BvrR/BvrS which controls the expression of various cell surface proteins (Omp 3 family) allowing Brucella to escape from the lysosomal pathway. After internalization virulent brucellae inhibit the fusion of their phagosomes with lysosomal compartments and modulate the phagosomal composition in order to evade intracellular degradation. The type IV secretion system encoded by the VirB operon is also a major virulence factor and plays an important role in intracellular trafficking.21 Effector molecules secreted by the type IV secretion machinery are responsible for maintaining the interactions of the Brucella-containing vacuole with the endoplasmic reticulum of the host cell until the replicative niche of Brucella is fully developed.22 Within this compartment, also called “brucellosome,” brucellae are able to survive and even replicate without restricting basic functions of the host

Molecular Detection of Foodborne Pathogens

cells. In the stationary phase intracellular bacteria still have to withstand severe nutrient deprivation and microaerobic conditions.23,24 Hence, a considerable degree of metabolic versatility is required for long-term survival and replication in this compartment. The majority of proteins regulated at this stage of infection are involved in primary metabolism of Brucella, confirming a ‘stealthy’ adaptation of the pathogen to the host cell environment.25

23.1.3  Brucellosis in Animals and Men Although many domestic animals are susceptible to various Brucella species, cattle, sheep, goats, and pigs are ­primarily affected. Animal brucellosis is characterized by abortion, weak newborn, retained placenta, orchitis, and impaired fertility.26 Hence, considerable economic damage is caused through the loss of progeny but also due to reduction in milk yield. Inapparent infections have been described in all susceptible animal species and spread between herds usually occurs by the introduction of these asymptomatic chronically infected animals. Brucellae are disseminated within herds of cattle and flocks via contamination of cowsheds and pastures through abortion or full-term parturition of infected carriers.27 The infection is mainly transmitted orally by licking aborted fetuses and genital discharges of an aborting animal or by feeding contaminated colostrum and milk. However, congenital (in utero) or perinatal infections may also occur. Especially in pigs venereal transmission is a significant route of spread since brucellae are often localized in the nongravid uterus. Four nomen species of the genus Brucella are pathogenic for humans, i.e., according to their impact on public health B. melitensis, B. abortus, B. suis, and rarely B. canis.1 Infrequent cases of brucellosis in humans caused by B. pinnipedialis or B. ceti have been described.28–30 Currently, only little is known about the pathogenicity of B. microti. Both the sources of infection and the species found may vary according to the geographical region. Brucella can be transmitted directly or indirectly from infected animals to humans and brucellosis usually is either an occupational or a foodborne infection. Direct transmission may involve the respiratory, conjunctival, and ­cutaneous route and requires close contact with infected animals. In endemic countries, brucellosis is recognized as an occupational disease and people engaged in the livestock industry, e.g., sheepherders, farmers, abattoir workers, and veterinarians, are often affected. Assistance in animal parturition is supposed to bear the highest risk of infection.31 The inhalation of aerosols containing viable brucellae is an important means of transmission to humans. The dose of infection by airborne transmission can be as low as ten bacteria and the attack rate of Brucella infections ranges between 30 and 100%. Therefore, brucellosis is the most frequent bacterial laboratory-acquired infection worldwide.32 However, indirect transmission remains the highest overall risk and mainly occurs through the consumption of unpasteurized milk or dairy products.31

Brucella

Human brucellosis is a disease of protean manifestations. The clinical onset of disease is insidious, and the incubation period is extremely variable ranging from weeks to months. Acute brucellosis is usually accompanied by bacteraemia and spreading of the organism to various organ systems mainly to reticuloendothelial tissues, i.e., liver, spleen, skeletal, and hematopoietic system. The capability of Brucella spp. to survive and replicate in the mononuclear phagocytic system is responsible for the high frequency of prolonged disease, complications, and relapses. Clinical manifestations may vary widely including osteoarticular, dermal, gastrointestinal, respiratory, cardiovascular, and neurologic disorders. Human brucellosis may mimic many other infectious as well as noninfectious diseases. Patients suffering from brucellosis usually present with nonspecific systemic signs and symptoms consistent with an influenza-like or septicemic illness, i.e., fever, fatigue, malaise, weight loss, headaches, arthralgia, myalgia, chills, and sweats. Brucella infections often develop as fever of unknown origin with an acute onset.33 Fever >38.5°C is the leading symptom in the majority of the patients. As this fever can wax and wane the disease is also called undulant fever.34 Osteoarticular manifestations, e.g., spondylitis, sacroiliitis, or arthritis are known to be the most frequent focal complications.35,36 Inadequate treatment may cause severe and debilitating chronic courses of infection and long-term sequelae. Patients can be plagued by serious life-threatening focal complications such as endocarditis and neurobrucellosis. Brucella endocarditis accounts for 80% of the fatal cases.37 The overall case fatality rate reported is low (less than 1%). Controlling acute illness and preventing complications and relapse are the most important tasks in the treatment of human brucellosis. To reach this goal prolonged chemotherapy using at least two synergistic antibiotic drugs is required. In the current guidelines for treating human brucellosis a combination of oral doxycycline 100 mg twice a day and rifampin 600–900 mg/day (15 mg/kg/day) in a single oral dose over a 6-week course is recommended.38 Nevertheless, the risk for a relapse remains high (about 10% of the patients appropriately treated).

23.1.4  Epidemiology of Brucellosis As brucellosis is a zoonotic disease transmitted from an animal reservoir to man, the global number of human cases can only be reduced by effective surveillance and control of animal brucellosis. Bovine brucellosis (B. abortus) has been successfully reduced in the last decades whereas the control of brucellosis in small ruminants (B. melitensis) has proven to be an intractable problem. Test and slaughter programs as well as the vaccination of livestock helped to eradicate brucellosis. However, the spill over of brucellosis from wildlife to livestock and the glob­ alization of animal trade promote the worldwide distribution of animal and human disease.2,39 Hence, despite being

319

controlled in many developed countries, brucellosis remains a major public health concern. Bovine brucellosis has been successfully eradicated in Canada, Japan, northern Europe and Australia. In the European Union, Sweden, Denmark, Finland, Germany, the United Kingdom (except for Northern Ireland), Austria, The Netherlands, Belgium, and Luxembourg are approved as “officially free from bovine and ovine/caprine brucellosis.”40 Norway and Switzerland are also considered to be brucellosis-free. In contrast, the situation is less favorable in Southern European countries.41 Ovine/caprine brucellosis endemically occurs in countries surrounding the Mediterranean Sea, especially along its northern and eastern shores stretching through Central Asia. Furthermore, it is highly prevalent in countries around the Arabian Gulf. Parts of Latin America, especially Mexico, Peru, and northern Argentina are also seriously affected. Last but not least, B. melitensis infections in sheep have been reported in Africa and India. B. melitensis is not known to be enzootic in the United States, Canada, northern Europe, Australia, New Zealand, or Southeast Asia which is why only sporadic incursions have been notified in these regions. Porcine brucellosis occurs in most areas in which pigs are kept. The human pathogenic biotypes, B. suis bv 1 and bv 3, are mainly isolated from wild boars, feral swine and domestic pigs in Southeast Asia, South America, in the south-eastern states of the USA and in Queensland, Australia.1,39 In Europe, porcine brucellosis is supposed to be restricted to biotype 2 which has only exceptionally been described as the causative agent of human brucellosis.42,43 The epidemiology of human brucellosis has significantly changed in the last decade for sanitary, socio-economic and political reasons. Currently, about half a million new cases are annually reported worldwide and prevalence rates exceed ten cases per 100,000 population in various endemic countries.2 Taking into account that less than 10% of the Brucella infections are supposed to be recognized and notified because of the unspecific clinical symptoms of the disease, human brucellosis represents a significant and probably neglected health problem.44,45 Although ovine/caprine brucellosis has a limited geographic distribution, B. melitensis is by far the main cause of clinically apparent disease in humans worldwide.46,47 In territories where ovine/caprine brucellosis is endemic and the density of the sheep and goat population is high, there is a significant relationship between the number of infected flocks and the incidence of human cases.48 B. abortus and B. suis infections are often associated with certain occupational groups, e.g., farm workers, veterinarians, and meatpacking employees whereas human B. melitensis infections frequently occur without occupational exposure.34 In the United States, the change of brucellosis from an occupational disease to a more common disease in the general population was accompanied by a shift from B. abortus infections most frequently reported before 1960 to B. suis infections predominating in the early 1970s and currently to an increased reporting of B. melitensis infections.49

320

In nonendemic countries, an association of brucellosis with the immigrant population has been described.50–52 Most cases have been imported and the disease was acquired either through international travel or infected food products from endemic areas.53 In the United States, 50–60% of the brucellosis cases are notified in California and Texas, mainly among Hispanics.44,54 Hispanic ethnicity, recent travel to endemic areas in Mexico and ingestion of nonpasteurized dairy products are the major risk factors for Brucella infections in the United States.44,49,52,54

23.1.5 Microbiological and Serological Methods in the Diagnosis of Brucellosis 23.1.5.1  Isolation and Identification All abortions in cattle, sheep, and goats have to be treated as suspected brucellosis and should be investigated. Whenever possible, Brucella should be isolated from potentially infected animals. Valuable samples can be vaginal secretions (swabs), uterine discharges, aborted fetuses (stomach contents, spleen, and lung), fetal membranes, udder secretions, milk, semen, and arthritis or hygroma fluids. From animal carcasses, tissues of the reticulo-endothelial system (lymph nodes and spleen), the late pregnant or early postparturient uterus, the udder, the testes or epididymes are preferred for culture. Brucella grows on most standard media, e.g., blood agar, chocolate agar, Trypticase soy agar (TSA) or serum-dextrose agar (SDA). 2–5% bovine or equine serum, which is needed for growth by various strains, is routinely added to the basal media. The inoculated agar plates should be incubated at 35–37°C in air supplemented with 5–10% CO2. Culturing the fastidious bacterium takes several days or even weeks. If samples of excreta or contaminated tissues are to be examined, selective media, containing amphotericin B, bacitracin, cycloheximide/natamycin, d-cycloserine, nalidixic acid, polymyxin B and vancomycin, can be used.55 As the number of Brucella organisms is likely to be low in milk, colostrum, some tissue samples and dairy products such as cheese, enrichment culture using liquid media may be helpful. Castañeda’s medium, a nonselective, biphasic medium, is usually recommended for the isolation of Brucella when enrichment culture is necessary.56 However, weekly subcultures on to solid selective medium for up to 6 weeks are necessary. After a 2–3 day incubation period punctuate, nonpigmented, and nonhemolytic Brucella colonies are visible. Colonies of smooth (S) brucellae are raised, convex, circular, translucent, and 0.5–1 mm in diameter. After sub- or prolonged (>4 days) culture, morphology as well as virulence, antigenic properties and phage-sensitivity tend to undergo variation. Smooth Brucella cultures dissociate to rough (R)-forms growing in less convex and more opaque colonies with a dull dry yellowish-white granular appearance. Suspected culture colonies can be confirmed by the slide agglutination test using undiluted polyvalent Brucella antiserum (anti-S-serum) mixed with a saline suspension of colonies.

Molecular Detection of Foodborne Pathogens

Brucella spp. are very small Gram-negative coccobacilli, which microscopically appear only faintly stained and look like “fine sand.” Oxidase and urease positive bacteria have to be suspected to be Brucella and have to be manipulated in a biological safety cabinet. The identification of Brucella species and biotypes is based on CO2 requirement, H2S production, urease activity, agglutination with monospecific sera (A and M), selective inhibition of growth on media containing dyes such as thionin or basic fuchsin, and phage typing.8,9 These procedures are time-consuming, hazardous, and subject to variable interpretation. Using commercially available biochemical tests such as the API 20 NE® (BioMerieux, Nürtingen, Germany) Brucella spp. may be misidentified as Moraxella phenylpyruvica.57 23.1.5.2  Serological Tests Because of the shortcomings of culture, serological tests are commonly used in control programs (ongoing diagnostic in order to detect positive animals in the international trade), eradication programs (reducing the prevalence of brucellosis in the population), and surveillance programs (continuous surveys of domestic animals for maintaining the disease free status of the population). However, the detection of high titers of specific antibodies in milk or serum samples only allows a tentative diagnosis. Cross-reactions with various bacteria, e.g., Yersinia enterocolitica O:9, Escherichia coli O:157, Francisella tularensis, Salmonella urbana group N, Vibrio cholerea, and Stenotrophomonas maltophilia, have been reported.58 Especially in the early stage of infection false-negative or only weak positive reactions may occur. Additionally, the sensitivity of various serological tests is low because they are based on secondary interactions such as the ability to agglutinate antigens or to fix complement. Particularly the quality of the antigen preparations is essential for consistent and reproducible results. Interpretation of test results is subjective reflecting the experience of the investigator. The buffered Brucella antigen tests, i.e., the Rose Bengal test (RBT) and the buffered plate agglutination test (BPAT), the complement fixation test (CFT), the enzyme-linked immunosorbent assay (ELISA) and the fluorescence polarization assay (FPA), are suitable for screening herds and individual animals (Table 23.1).59,60 RBT and CFT are prescribed for the screening of flocks and individual sheep or goats.61 However, there is no single serological test which might be appropriate for all epidemiological situations and due to the low specificity of serological tests, a confirmatory test is always recommended to avoid false positive results.62,63 Samples that reacted positive in a screening test are retested using CFT and ELISA, both of which are suitable for screening and confirmation. Because of its low sensitivity and specificity as compared to other serological tests, the serum agglutination test (SAT) does not meet the requirements of a screening test in international trade and is neither recommended for use in cattle nor in small ruminants.

321

Brucella

Table 23.1 Mean Sensitivity and Specificity Values of Various Serological Tests Routinely Used in the Diagnosis of Bovine Brucellosis Test Culture RBT BPAT CFT iELISA cELISA FPA Milk ring test Milk iELISA Milk FPA

Sensitivity (%)

Specificity (%)

46 81 95 89 96 98 98 90 98 100

100 86 98 84 94 91 99 75 96 96

Data compiled from various data sets. Adapted from Gall, D. and Nielsen, K., Rev. Sci. Technol. Off. Int. Epiz., 23, 989, 2004.

The milk ring test and the indirect ELISA which can be performed on bulk milk samples are effective for screening and monitoring dairy cattle for brucellosis. Both methods are efficient to identify herds with infected animals, even if the prevalence of brucellosis within the herd is low. However, the milk ring test is less reliable in large herds and should then be modified by either increasing the sample volume tested or by taking segmented samples. The milk ring test is carried out at intervals of 4–6 months and all animals in a herd have to be serologically tested when a positive test result is obtained. Although the milk ring test is certainly more efficient in detecting infected herds than market or slaughterhouse testing, the latter strategy might be more cost effective with decreasing incidence and prevalence of brucellosis. However, trace-back to the herd of origin must be possible at any time in the food production chain. For the serological screening of goats and sheep, there is no test equivalent to the bulk milk testing in cattle. The brucellin allergic skin test which is based on cellular immunity can be used as a screening or as a confirmatory test in unvaccinated herds and flocks. The test is carried out by intradermal injection of a purified (free of sLPS), standardized antigen preparation (e.g., brucellin INRA) either in the cervical or caudal fold regions and 72 hours after injection a swelling ≥2 mm is considered to be positive. Since the specificity of the brucellin skin test is very high (95–100%) positive results may help to differentiate false positive serological reactions caused by infections with cross-reacting bacteria from Brucella infected animals, especially in brucellosisfree areas.64,65 Conventional serological tests for antibody detection. RBT and BPAT: The RBT has been developed in the United States as a rapid screening test in field surveillance of domestic cattle especially in endemic areas. The RBT is a card test using a B. abortus strain 99 (Weybridge) or B. abortus strain 1119-3 (USDA) antigen suspension (8%) stained with Rose Bengal dye buffered to pH 3.65±0.05.66 Acidification loosens

the bond between the antigen and non-specific agglutinins leading to an improved specificity and sensitivity of the RBT and the BPAT (pH 3.7±0.03) in comparison with SAT.67,68 CFT: CFT is recommended as confirmatory test in the diagnosis of brucellosis because it is considered to be the most sensitive and specific conventional serological method.69,70 Especially in the late chronic stage of disease or after vaccination when SAT results are either negative or inconclusive, CFT has proven to be the diagnostic tool of choice.71,72 Primary binding assays for antibody detection. ELISA: A great variety of antigens has been used for indirect enzymelinked immunosorbent assay (iELISA) including whole cells, sLPS, native hapten polysaccharide, salt-extracted proteins, outer membrane proteins and cytoplasmic proteins.58 However, most ELISAs are based on bacterial extracts including high amounts of LPS and false-positive reactions can occur after exposure to cross-reacting bacteria or after vaccination. Competitive enzyme immunoassays (cELISAs) are less prone to cross-reacting antibodies than conventional tests or iELISAs. They are also capable of differentiating vaccination antibodies from antibodies after natural infection.73,74 Because of their additional high sensitivity cELISAs can be used as confirmatory and screening tests. FPA: The principle of fluorescence polarization is that light will be depolarized more intensively by a small fast rotating molecule than by a larger molecule. Since every molecule in a solution rotates randomly at a rate inversely proportional to its size, the rotation rate of a small antigenic molecule, measured using a fluorescent label and polarized light, will change after antibodies have been attached. The FPA is capable of distinguishing vaccinated animals from naturally infected animals comparable to the cELISA.75 The specificity of the FPA in vaccinated animals rises with the time passed after inoculation.76 Furthermore, the FPA may help to eliminate unspecific reactions caused by crossreacting bacteria. The FPA can be performed within a few minutes and requires minimal preparation. Under field conditions even whole blood samples can be analyzed.77

23.1.6  Molecular Diagnostic Techniques 23.1.6.1  PCR Numerous PCR techniques have been developed for the identification of Brucella.78 PCR turned out to be a suitable tool for the detection of Brucella in various matrices including bovine blood and milk,79–82 organs of naturally infected cattle80,83 as well as milk and cheese of goats or sheep.84–86 PCR methods are increasingly used in the diagnosis of brucellosis since they can be more sensitive than blood culture and more specific than serological tests.8 In addition, the work with DNA as opposed to highly infectious live cultures decreases the risk of laboratory-acquired infections. However, the results of the few studies conducted on molecular detection of Brucella in animals and food products are highly controversial. Romero et al. reported that serological methods surpassed PCR detection in bovine milk samples.82 O’Leary et al. also proved that bacteriological

322

methods identified a higher number of Brucella infected cows than PCR.80 In contrast, PCR detection exceeded serological and bacteriological methods in experimentally and naturally infected goats.87,88 Furthermore, Hamdy and Amin reported that PCR detection has proven to be more successful in bovine milk samples as compared to microbiological methods whereas more positive cases were detected in sheep samples by bacteriological techniques.87 For large-scale field screening and to exclude the contamination of food products, the simple identification of Brucella by genus-specific PCR is adequate. A lot of target sequences have been used for detection of the genus Brucella, e.g., the 16S rRNA,89,90 the 16S–23S internal transcribed spacer region (ITS),91 a gene coding for an outer membrane protein (omp2),79,88,92 and the insertion element IS711.93,94 However, the majority of genus-specific PCR assays target the bcsp31 gene.95 The amplification product consists of 223 bp within a gene encoding a 31-kDa immunogenic outer membrane protein conserved among Brucella spp. After inoculation and haematogenous dissemination, Brucella organisms were present in all organs of infected cows and the bcsp31 PCR specifically detected Brucella DNA in the uterus, udder, spleen, lymph nodes, liver, and kidney after enrichment.83 For epidemiological trace back or species-specific eradication programs differential PCR assays are needed. The multiplex AMOS-PCR using a five primer cocktail was a first approach towards the differentiation of four Brucella species, i.e., B. abortus, B. melitensis, B. ovis, and B. suis.96 One primer anneals to the insertion sequence IS711 which appears in several copies all over the genome but distribution and number differ in Brucella species. The other four primers hybridize to species-specific regions downstream this insertion sequence leading to variable amplicon sizes. Recently, a conventional multiplex PCR assay suitable for the identification of all Brucella species and the vaccine strains, B. abortus RB51, B. abortus S19, and B. melitensis Rev1, has been developed.97 Real-time PCR constitutes a further technological improvement for the molecular identification of the genus Brucella and the differentiation of its species. Especially in the context of biological warfare and agroterrorism real-time PCR will allow a more rapid high-throughput screening of samples.98,99 Real-time PCR has proven to be a valuable tool when culture fails or serological results are inconclusive, and in addition, a test result can be available within a few hours. The species-specific real-time PCRs established by Redkar et al.,100 Newby et al.,101and Probert et al.102 were described as rapid, sensitive, and specific diagnostic tools with a low risk of cross-contamination and the potential of automation. SNP (single nucleotide polymorphishm)-based differentiation of Brucella clades incorporated into real-time PCR assays may also provide a rapid method for the identification of Brucella species.103 Various studies of genus-specific real-time PCR assays applied to human and animal specimens have been published whereas studies on the detection of Brucella DNA in food products using real-time PCR are rather scarce. A real-time PCR targeting the bcsp31 gene appears to be suitable for screening whenever Brucella is suspected because

Molecular Detection of Foodborne Pathogens

false negative results caused by rare species and biotypes may be avoided by a genus-specific assay. Consecutively, a species-specific assay should be applied keeping the epidemiological background in mind in order to confirm the diagnosis with a second gene target.99 The use of more than one marker-based PCR has proven to increase sensitivity and specificity in the molecular diagnosis of brucellosis in field animals and seems to be superior to ELISA at least when screening blood samples.104 Since PCR reactions may be inhibited in food samples, an internal amplification control should be included.99 Dilution of the samples may also reduce inhibitory effects of the sample matrix but the generally small amount of DNA in environmental and clinical samples is challenging even for assays with a very low detection limit. Approximately five bacteria per reaction can be detected using well established Brucella real-time PCR assays. Testing several replicates of the purified DNA in parallel may increase the probability of a correct result. 23.1.6.2  Molecular Typing Methods Molecular epidemiology significantly contributes to the analysis and understanding of infections caused by pathogenic bacteria.105 In epidemiological studies, molecular typing methods can be used for trace-back analysis which may help to identify the origin of an infection and the way of its spreading. Hence, appropriate countermeasures can be implemented in a timely manner and further spread of the disease will be prevented. Fast and accurate typing procedures are therefore crucial for eradication and control of brucellosis. In geographical regions where specific species or biovars may predominate, e.g., B. melitensis in the Mediterranean Basin, the classical biotyping methods cannot be used effectively for trace-back investigations. Furthermore, Brucella spp. have proven challenging to differentiate using molecular techniques due to the high degree of genetic homology (>90%) among species demonstrated by DNA–DNA hybridization,14 due to identical 16S rRNA sequences among all Brucella species,106 and because more than 90% of all genes share >98% sequence identity.107 Nevertheless, various methods have been established for molecular subtyping of Brucella strains, e.g., AP-PCR (arbitrary primed-PCR),108 ERIC-PCR (enterobacterial repetitive intergenic consensus sequence-PCR), REPPCR (repetitive intergenic palindromic sequence-PCR),109,110 RAPD-PCR (random amplified polymorphic DNA-PCR),111 PCR-RFLP (PCR-restriction fragment length polymorphism) of different genetic loci,112 AFLP (amplified fragment length polymorphism),113 SNP114,115 and MLST (multi locus sequence typing).116 However, most of these tests lack reproducibility, show a limited capability to differentiate single strains, are more demanding than simple PCR typing methods or are not appropriate for routine typing. Recently, the genomes of B. melitensis 16M,117 B. suis 1330118 and B. abortus strain 9-941107 have been sequenced completely. DNA sequence variability required for subtyping or epidemiological trace-back of Brucella strains can now be deduced by direct comparison of genome sequences. Hence, tandem repeat loci predicted to display size polymorphism

323

Brucella

could be identified. Variable number tandem repeats (VNTRs) which are commonly used for DNA fingerprinting in forensic applications also exist in bacterial genomes and seem to be highly discriminatory markers, even when the pathogens investigated belong to monomorphic species such as Brucella.119–121 Tandem repeats are composed of perfect or imperfect copies of an elementary unit and various alleles can be observed in different bacterial strains within a species. Fingerprints resulting from analysis of multiple loci can be highly discriminating or even unique. Multiple locus variable number tandem repeats analysis (MLVA) has proven to be highly discriminatory among unrelated Brucella isolates that could not be differentiated by classical microbiological methods. The hypervariability found by MLVA is in contrast to the well-known genetic homogeneity of the genus. However, tandem repeats generally mutate at different rates.122 Some loci cluster Brucella isolates in accordance with the identified species, presumably because they mutate slowly and have a relatively low homoplasy rate. Others have a higher discriminatory index and Brucella isolates originating from a restricted geographical area can still be discriminated indicating the potential of VNTRs as an epidemiological tool.119 The hypermutability of VNTR loci has already been exploited for Brucella strain typing in large panels of animal isolates.120–124 Based on simple PCR techniques, Brucella-MLVA is accessible to a wide range of users. The alleles (PCR amplicons) can be analyzed by simple agarose gel electrophoresis or automatic high-throughput procedures. Standardization and quality control are easy to achieve. Especially in an outbreak setting including a great number of cases high-throughput screening is possible without manipulation of the living agent. Last but not least, the MLVA data can be easily coded and exchanged by the repeat copy numbers for each locus and strain. Currently, MLVA is the most suitable molecular method for subtyping brucellae which fulfils all the recommended performance criteria of a typing assay, i.e., typeability, reproducibility, stability, discriminatory power, concordance with other typing techniques and epidemiological concordance.123

fresh milk and tissue, and also from paraffin-embedded tissue samples.80,126 Queipo-Ortuno et al. evaluated seven commercially available DNA extraction kits using human serum samples and obtained the best results with the UltraCleanTM DNA BloodSpin Kit (MO BIO Laboratories Inc., Carlsbad, CA),127 which was also successfully used by Navarro et al.128 However, comprehensive evaluation studies on DNA extraction methods in animal and food samples are still missing.

23.2.2  Detection Procedures 23.2.2.1  Multiplex PCR A previously described AMOS-PCR is highly effective for differentiation of B. abortus, B. melitensis, B. ovis and B. suis96 (Table 23.2). This PCR employs one primer (IS711) for binding with the insertion sequence IS711, which is present in all Brucella species, and four primers for recognizing the speciesspecific regions downstream of this insertion sequence, leading to formation of variable sizes of amplicons from the four Brucella species, and facilitating their identification (Figure 23.1). Table 23.2 Primers for AMOS-PCR Primer BA BM BS BO IS711

Sequence 5´-GAC GAA CGG AAT TTT TCC AAT CCC-3´ 5´-AAA TCG CGT CCT TGC TGG TCT GA-3´ 5´-GCG CGG TTT TCT GAA GGT TCA GG-3´ 5´-CGG GTT CTG GCA CCA TCG TCG-3´ 5´-TGC CGA TCA CTT AAG GGC CTT CAT-3´

23.2  Methods

B. melitensis

B. suis

Both specificity and sensitivity of the most commonly used bcsp31 PCR have proven to be high at least in colony material.99,125 The limit of detection can be lower than 20 fg of DNA when determined on the basis of purified DNA obtained from colonies. In contrast, the low number of bacteria usually present in clinical samples may be a limiting factor for real-time PCR assays. Due to the small number of brucellae in clinical specimens, special methods for DNA preparation are required to reduce inhibitory effects caused by matrix components and to concentrate the DNA. Blood, milk, diary products, tissue, semen, and vaginal swabs can be valuable specimens for direct detection of Brucella DNA. Commercial kits such as the QIAampTM DNA Mini Kit (Qiagen Inc., Valencia, CA) can be used to extract DNA from

B. abortus

23.2.1  Sample Collection and Preparation

Figure 23.1  Identification and differentiation of B. abortus, B. melitensis, and B. suis using AMOS-PCR based on the repetitive DNA sequence IS711. (From Bricker, B.J. and Halling, S.M., J. Clin. Microbiol., 11, 2660, 1994.)

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Molecular Detection of Foodborne Pathogens

Table 23.3 Primers for Brucella Multiplex PCR Primers BMEI0998f BMEI0997r BMEI0535f BMEI0536r BMEII0843f BMEII0844r BMEI1436f BMEI1435r BMEII0428f BMEII0428r BR0953f BR0953r BMEI0752f BMEI0752r BMEII0987f BMEII0987r Bmispec f Bmispec r

Sequence

Amplicon Size (bp)

5´-ATC CTA TTG CCC CGA TAA GG-3´ 5´-GCT TCG CAT TTT CAC TGT AGC-3´ 5´-GCG CAT TCT TCG GTT ATG AA-3´ 5´-CGC AGG CGA AAA CAG CTA TAA-3´ 5´-TTT ACA CAG GCA ATC CAG CA-3´ 5´-GCG TCC AGT TGT TGT TGA TG-3´ 5´-ACG CAG ACG ACC TTC GGT AT-3´ 5´-TTT ATC CAT CGC CCT GTC AC-3´ 5´-GCC GCT ATT ATG TGG ACT GG-3´ 5´-AAT GAC TTC ACG GTC GTT CG-3´ 5´-GGA ACA CTA CGC CAC CTT GT-3´ 5´-GAT GGA GCA AAC GCT GAA G-3´ 5´-CAG GCA AAC CCT CAG AAG C-3´ 5´-GAT GTG GTA ACG CAC ACC AA-3´ 5´-CGC AGA CAG TGA CCA TCA AA-3´ 5´-GTA TTC AGC CCC CGT TAC CT-3´ 5´-AGA TAC TGG AAC ATA GCC CG-3´ 5´-ATA CTC AGG CAG GAT ACC GC-3´

1,682 450 (1,320 in Brucella strains isolated from marine mammals) 1,071 794 587 272 218 152 510

BMEI and BMEII numbers designate loci in the B. melitensis genome; BR numbers designate loci in the B. suis genome; f, forward; r, reverse.

Table 23.4 Multiplex PCR Cycling Program Cycles 1 25

23.2.2.2  Real-Time PCR Real-time PCR assays targeting bcsp31 described by Probert et al. and Al Dahouk et al. can be recommended for the detection of Brucella for screening purposes (Tables 23.1 through 23.6).99,102 Assays for the species-specific identification of B. melitensis described by Redkar et al. and Probert et al. are also very sensitive (Table 23.8).99,100,102 Assays designed for the detection of B. abortus by Redkar et al., Probert et al., and Newby et al. only detected biotypes 1, 2, and 4, because of the more distant relatedness of other biotypes (Table 23.7).100–102 A real-time PCR assay specific for B. suis biotype 1 was also established by Redkar et al.100

B. microti

B. ceti

B. pinnipedialis

B. neotomae

To identify all Brucella species and the vaccine strains, B. abortus RB51, B. abortus S19, and B. melitensis Rev1, another multiplex PCR assay97 can be applied. Use of primers and PCR conditions shown in Tables 23.3 and 23.4 will result in the amplification of distinct bands from all Brucella species (Figure 23.2).97

B. canis

1

B. ovis

15 min 30 sec 90 sec 120 sec 10 min

B. melitensis

Time

95 94 58 72 72

B. suis

Temperature (°C)

B. abortus

Initial denaturation Denaturation Annealing Extension Last extension

Figure 23.2  Identification and differentiation of all Brucella species by a multi-locus multiplex PCR. (From García-Yoldi, D. et al., Clin. Chem., 52, 779, 2006.)

Hybridization probes technology offers melting curve analysis which provides typical signals for specific amplification products. 5′-exonuclease assays are very useful for large scale screening as they can be performed on platforms including 96 or 384 tests in parallel. Since real-time PCR assays for the detection of biologic agents can be easily transferred from one technical platform to another,129 diagnostic laboratories are able to choose between hybridization probes technology and 5′-exonuclease assays to meet

325

Brucella

Table 23.5 Real-Time PCR Assay Targeting bcsp31 for the Specific Detection of the Genus Brucella A. PCR mixture makeup Reagent

Stock Concentration (pmol/µl=µM)

Brucella spp. (bcsp31)

Water 25 mM MgCl2 10 × concentrated Reaction mix

LightCycler FastStart DNA Master Hybridization Probes Cat. No. 2 239 272

Primers and probes (5´-3´) B4 B5 Bru FL Bru LC Lambda F Lambda R Lambda FL Lambda LC Phage lambda DNA* Sample DNA

TGGCTCGGTTGCCAATATCAA CGCGCTTGCCTTTCAGGTCTG AGGCAACGTCTGACTGCGTAAAGCC LC Red 640-ACTCCAGAGCGCCCGACTTGATCG ATGCCACGTAAGCGAAACA GCATAAACGAAGCAGTCGAGT GGTGCCGTTCACTTCCCGAATAAC X LC Red 705-CGGATATTTTTGATCTGACCGAAGCG p Plasmid DNA

µl Per Reaction

Final Concentration (in 20 µl)

9.8 1.2

2.5 mM

2

20 pmol/µl 20 pmol/µl 8 µM 8 µM 20 pmol/µl 20 pmol/µl 8 µM 8 µM

0.5 0.5 0.5 0.5 0.5 0.5 0.5 0.5 1 2

0.5 µM 0.5 µM 0.2 µM 0.2 µM 0.5 µM 0.5 µM 0.15 µM 0.15 µM

Temperature (°C)

Time

Slope (°C/s)

Acquisition Mode

95

10 min

20

None

95 55 72 95 55 95 40

10 sec 10 sec 12 sec 0 30 sec 0 30 sec

20 20 20 20 20 0.1 20

None Single None None None Continuous None

B. PCR Parameter Parameter Activation of Fast Start Taq DNA polymerase Amplification (45 cycles)

Melting curve analysis Cooling

The assay specifically detects all brucellae but no other organisms with a detection limit of 16 fg DNA in buffer. *Inclusion of an internal amplification control is especially useful for clinical samples (Al Dahouk, S. et al., Clin. Chem. Lab. Med., 45, 1464, 2007).

their demands. Ready to use protocols for the detection of Brucella and the identification of the most relevant species by means of real-time PCR assays are given in the Tables 23.5 through 23.8.

23.3 Conclusions and Future Perspectives In the diagnosis of brucellosis time-consuming culture and phenotypic characterization of the isolate is the “gold standard.” The low yield of Brucella cultures, however, often results in diagnostic delay and the late initiation of appropriate countermeasures. Serology is a more effective means of diagnostic, although cross-reactivity in the setting of other bacterial infections is still a major problem. PCR has proven to be a valuable tool when culture fails or serological results are inconclusive, and in addition, a test result can be available within a few hours. PCR assays targeting more than one gene of the Brucella genome may enhance the likelihood of detection. Genus-specific primers and probes

should always be included in a diagnostic scheme to detect infrequently isolated Brucella species and atypical strains. However, a positive result in a genus-specific real-time PCR has to be specified by a species-specific assay. Serological tests should be performed in parallel to further consolidate the diagnosis as the number of constantly seronegative animals is low. There has been controversial discussion about the preferred clinical specimen, the best method of DNA extraction, and the PCR assays with the lowest limit of detection. Most results were not reproducible in different laboratories which demonstrate the complexity of PCR procedures including DNA extraction from clinical specimens. PCR assays targeting different genes may help to exclude false positive results due to contamination. In order to determine inhibitory effects of the matrix an internal amplification control should be integrated in all assays. Since practical experiences may vary between laboratories concerning optimal DNA preparation and PCR assays, results obtained with molecular methods generally have to be evaluated in

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Molecular Detection of Foodborne Pathogens

Table 23.6 Real-time 5′-Exonuclease Assay Targeting bcsp31 for Large Scale Screening of the Genus Brucella A. PCR Mixture Makeup Reagent

Stock Concentration (pmol/µl=µM)

Brucella spp. (bcsp31)

5.5

Water Reaction buffer

µl Per Reaction

TaqMan Universal MasterMix (uMM) Applied Biosystems Cat. No. 4304447

2×

Final Concentration (in 25 µl)  

12.5

1 ×

Primers and probes (5´-3´) Brucella spp fw GCTCGGTTGCCAATATCAATGC

10 µM

0.75

0.3 µM

Brucella spp rev

GGGTAAAGCGTCGCCAGAAG

10 µM

0.75

0.3 µM

Brucella spp. Taq

6FAM-AAATCTTCCACCTTGCCCTTGCCATCA-DB

10 µM

0.5

0.2 µM

Sample DNA

5 B. PCR parameter Temperature (°C)

Time

Cycle 1

Decontamination

50

2 min

Initial denaturation

95

10 min

1

Denaturation

95

15 sec

45

Annealing

57

1 min

The assay specifically detects all brucellae but no other organisms with a detection limit of 16 fg DNA in buffer. Source:  Probert, W.S. et al., J. Clin. Microbiol., 42, 1290, 2004.

Table 23.7 Real-Time PCR Assay Targeting alkB/IS711 for the Specific Detection of B. abortus Designed as 5′-Exonuclease Assay for Large Scale Screening A. PCR Mixture Makeup Reagent

Brucella abortus alkB/IS711

Stock Concentration (pmol/µl=µM)

Water Reaction buffer

TaqMan Universal MasterMix (uMM) Applied Biosystems

2×

µl Per Reaction

Final Concentration (in 25 µl)

5.5

 

12.5

1 ×

Primers and probes (5´-3´) B. abortus fw GCGGCTTTTCTATCACGGTATTC

10 µM

0.75

0.3 µM

B. abortus rev

CATGCGCTATGATCTGGTTACG

10 µM

0.75

0.3 µM

B. abortus Taq

6FAM-CGCTCATGCTCGCCAGACTTCAATG-DB

10 µM

0.5

0.2 µM

Sample DNA

5 B. PCR Parameter Temperature (°C)

Time

Cycle

Decontamination

50

2 min

1

Initial denaturation

95

10 min

1

Denaturation

95

15 sec

45

Annealing

57

1 min

This assay specifically detects B. abortus bv 1, 2, and 4, but other biotypes cannot be detected reliably. The detection limit is 18 fg DNA in buffer. Source:  Probert, W.S. et al., J. Clin. Microbiol., 42, 1290, 2004.

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Brucella

Table 23.8 Real-Time PCR Assay Targeting BMEI1162/IS711 for the Specific Detection of Brucella melitensis Designed as 5′-Exonuclease Assay for Large Scale Screening A. PCR Mixture Makeup Reagent

Stock Concentration (pmol/µl=µM)

Brucella Melitensis BMEI1162/IS711

Water Reaction buffer

TaqMan Universal MasterMix (uMM) Applied Biosystems

2×

Primers and probes (5´-3´) B. melitensis fw AACAAGCGGCACCCCTAAAA

µl Per Reaction

Final Concentration (in 25 µl)

5,5

 

12.5

1 ×

10 µM

0.75

0.3 µM

B. melitensis rev

CATGCGCTATGATCTGGTTACG

10 µM

0.75

0.3 µM

B. melitensis Taq

6FAM-CAGGAGTGTTTCGGCTCAGAATAATCCACA-DB

10 µM

0.5

0.2 µM

Sample DNA

5 B. PCR Parameter Temperature (°C)

Time

Cycle

Decontamination

50

2 min

1

Initial denaturation

95

10 min

1

Denaturation

95

15 sec

45

Annealing

57

1 min

This assay specifically detects all B. melitensis isolates but no other organisms with a detection limit of 16 fg DNA in buffer. Source:  Probert, W.S. et al., J. Clin. Microbiol., 42, 1290, 2004.

the context of other diagnostic tests. Currently, it seems to be unlikely that conventional or real-time PCR will supersede microbiological methods for detection of Brucella in clinical samples. For instance, O’Leary et al. obtained positive real-time PCR results only in a small proportion of culture-positive milk (44%) and supramammary lymph tissue samples (75%) in cattle naturally infected with B. abortus.80 Debeaumont et al. tested serum samples of brucellosis patients for their bacterial DNA load by a quantitative bcsp31-based real-time PCR.130 Approximately 25–650 genomic copies per 5 µl of DNA extract were found. QueipoOrtuno et al. were able to detect one bacterial cell in a serial dilution spiked with B. abortus B19 in 200 µl serum.131 Hence, the small sample volume (2–5 µl) used in real-time PCR systems may also be a serious disadvantage. The main reasons why PCR results in the diagnosis of brucellosis have always been variable when compared to serology and bacteriology are (i) the stage of infection which may influence the number and location of bacteria, (ii) the presence of large amounts of host genomic DNA that may have inhibitory effects on the PCR assay, (iii) the method of DNA extraction which may be crucial for the detection of the bacterium by molecular methods. Most real-time PCR assays developed for the detection of Brucella can detect less than ten bacteria per reaction, which is close to the technical limit of this method. Additionally, a variety of DNA purification methods has been established to

eliminate inhibitory components of the matrix. Limitations are the small number of brucellae in clinical specimens of infected animals and the small amount of DNA used in PCR assays. Due to these biological and technical limitations, no significant advantage of PCR methods compared to standard serological and bacteriological methods could have been demonstrated so far.

References



1. Godfroid, J. et al. From the discovery of the Malta fever’s agent to the discovery of a marine mammal reservoir, brucellosis has continuously been a re-emerging zoonosis. Vet. Res., 36, 13, 2005. 2. Pappas, G. et al. The new global map of human brucellosis. Lancet Infect. Dis., 6, 91, 2006. 3. Greenfield, R.A. et al. Bacterial pathogens as biological weapons and agents of bioterrorism. Am. J. Med. Sci., 323, 299, 2002. 4. Christopher, G.W. et al. History of U.S. military contributions to the study of bacterial zoonoses. Mil. Med., 170, 39, 2005. 5. Corbel, M.J. Microbiology of the genus Brucella. In Brucellosis: Clinical and Laboratory Aspects, p. 53. Young, E.J. and Corbel, M.J. (Eds.). CRC Press, FL, 1989. 6. Jumas-Bilak, E. et al. Unconventional genomic organization in the alpha subgroup of the Proteobacteria. J. Bacteriol., 180, 2749, 1998. 7. Moreno, E. et al. Brucella abortus 16S rRNA and lipid A reveal a phylogenetic relationship with members of the alpha-2 subdivision of the class Proteobacteria. J. Bacteriol., 172, 3569, 1990.

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8. Al Dahouk, S. et al. Laboratory-based diagnosis of brucellosis – a review of the literature. Part I: techniques for direct detection and identification of Brucella spp. Clin. Lab., 49, 487, 2003. 9. Alton, G.G. et al. Techniques for the Brucellosis Laboratory. Institut National de la Recherche Agronomique, Paris, 1988. 10. Corbel, M.J. and Brinley-Morgan, W.J. Genus Brucella. In Bergey’s Manual of Systematic Bacteriology, vol. 1, p. 370. Krieg, N.R. and Holt, J.G. (Eds.). Williams & Wilkins, Baltimore, MD, 1984. 11. Cloeckaert, A. et al. Classification of Brucella spp. isolated from marine mammals by DNA polymorphism at the omp2 locus. Microbes Infect., 3, 729, 2001. 12. Foster, G. et al. Brucella ceti sp. nov. and Brucella pinnipedialis sp. nov. for Brucella strains with cetaceans and seals as their preferred hosts. Int. J. Syst. Evol. Microbiol., 57, 2688, 2007. 13. Scholz, H.C. et al. Brucella microti sp. nov., isolated from the common vole Microtus arvalis. Int. J. Syst. Evol. Microbiol., 58, 375, 2008. 14. Verger, J.M. et al. Brucella, a monospecific genus as shown by deoxyribonucleic acid hybridization. Int. J. Syst. Bacteriol., 35, 292, 1985. 15. Osterman, B. and Moriyón, I. International Committee on Systematics of Prokaryotes Subcommittee on the Taxonomy of Brucella. Minutes of the meeting, 17 September 2003, Pamplona, Spain. Int. J. Syst. Evol. Microbiol., 56, 1173, 2006. 16. Moreno, E. and Moriyón, I. Brucella melitensis: a nasty bug with hidden credentials for virulence. Proc. Natl. Acad. Sci. USA, 99, 1, 2002. 17. Lapaque, N. et al. Brucella lipopolysaccharide acts as a virulence factor. Curr. Opin. Microbiol., 8, 60, 2005. 18. Porte, F. et al. Role of the Brucella suis lipopolysaccharide O antigen in phagosomal genesis and in inhibition of phagosome-lysosome fusion in murine macrophages. Infect. Immun., 71, 1481, 2003. 19. Forestier, C. et al. Brucella abortus lipopolysaccharide in murine peritoneal macrophages acts as a down-regulator of T cell activation. J. Immunol., 165, 5202, 2000. 20. Roop II, R.M. et al. Adaptation of the brucellae to their intracellular niche. Mol. Microbiol., 52, 621, 2004. 21. Boschiroli, M.L. et al. The Brucella suis virB operon is induced intracellularly in macrophages. Proc. Natl. Acad. Sci. USA, 99, 1544, 2002. 22. Celli, J. and Gorvel, J.P. Organelle robbery: Brucella interactions with the endoplasmic reticulum. Curr. Opin. Microbiol., 7, 1, 2004. 23. Köhler, S. et al. The analysis of the intramacrophagic virulome of Brucella suis deciphers the environment encountered by the pathogen inside the macrophage host cell. Proc. Natl. Acad. Sci. USA, 99, 15711, 2002. 24. Köhler, S. et al. What is the nature of the replicative niche of a stealthy bug named Brucella? Trends Microbiol., 11, 215, 2003. 25. Al Dahouk, S. et al. Quantitative analysis of the intramacrophagic proteome of the pathogen Brucella suis reveals metabolic adaptation to the late stage of cellular infection. Proteomics, 8, 3862, 2008. 26. Miller, M.A. and Paige, J.C. Other food borne infections. Vet. Clin. North Am. Food Anim. Pract., 14, 71–89, 1998. 27. Bercovich, Z. Maintenance of Brucella abortus-free herds: a review with emphasis on the epidemiology and the problems in diagnosing brucellosis in areas of low prevalence. Vet. Quart., 20, 81, 1998.

Molecular Detection of Foodborne Pathogens 28. Brew, S.D. et al. Human exposure to Brucella recovered from a sea mammal. Vet. Rec., 144, 483, 1999. 29. McDonald, W.L. et al. Characterization of a Brucella sp. strain as marine-mammal type despite isolation from a patient with spinal osteomyelitis in New Zealand. J. Clin. Microbiol., 44, 4363, 2006. 30. Sohn, A.H. et al. Human neurobrucellosis with intracerebral granuloma caused by a marine mammal Brucella sp. Emerg. Infect. Dis., 9, 485, 2003. 31. Cooper, C.W. Risk factors in transmission of brucellosis from animals to humans in Saudi Arabia. Trans. R. Soc. Trop. Med. Hyg., 86, 206, 1992. 32. Yagupsky, P. and Baron, E.J. Laboratory exposures to Brucellae and implications for bioterrorism. Emerg. Infect. Dis., 8, 1180, 2005. 33. Saltoglu, N. et al. Fever of unknown origin in Turkey: evaluation of 87 cases during a nine-year-period of study. J. Infect., 48, 81, 2004. 34. Young, E.J. An overview of human brucellosis. Clin. Infect. Dis., 21, 283, 1995. 35. Colmenero, J.D. et al. Complications associated with Brucella melitensis infection: a study of 530 cases. Medicine (Baltimore), 75, 195, 1996. 36. Gür, A. et al. Complications of brucellosis in different age groups: a study of 283 cases in southeastern Anatolia of Turkey. Yonsei Med. J., 44, 33, 2003. 37. Peery, T.M. and Belter, L.F. Brucellosis and heart disease. II. Fatal brucellosis: a review of the literature and report of new cases. Am. J. Pathol., 36, 673, 1960. 38. Food andAgriculture Organization-World Health Organization, Joint FAO/WHO Expert Committee on Brucellosis (sixth report). In WHO Technical Report Series, No. 740, p. 56. World Health Organization, Geneva, 1986. 39. Godfroid, J. Brucellosis in wildlife. Rev. Sci. Tech., 21, 277, 2002. 40. Godfroid, J. and Käsbohrer, A. Brucellosis in the European Union and Norway at the turn of the twenty-first century. Vet. Microbiol., 90, 135, 2002. 41. Taleski, V. et al. An overview of the epidemiology and epizo­ otology of brucellosis in selected countries of Central and Southeast Europe. Vet. Microbiol., 90, 147, 2002. 42. Paton, N.I. et al. Visceral abscesses due to Brucella suis infection in a retired pig farmer. Clin. Infect. Dis., 32, 129, 2001. 43. Teyssou, R. et al. About a case of human brucellosis due to Brucella suis biovar 2. Mèd. Mal. Infect., 19, 160, 1989. 44. Chomel, B.B. et al. Changing trends in the epidemiology of human brucellosis in California from 1973 to 1992: A shift toward foodborne transmission. J. Infect. Dis., 170, 1216, 1994. 45. Wise, R.I. Brucellosis in the United States: Past, present, and future. JAMA, 244, 2318, 1980. 46. Corbel, M.J. Brucellosis: An overview. Emerg. Infect. Dis., 3, 213, 1997. 47. Pappas, G. et al. Medical Progress – Brucellosis. N. Engl. J. Med., 352, 2325, 2005. 48. De Massis, F. et al. Correlation between animal and human brucellosis in Italy during the period 1997–2002. Clin. Microbiol. Infect., 11, 632, 2005. 49. Fosgate, G.T. et al. Time-space clustering of human brucellosis, California, 1973–1992. Emerg. Infect. Dis., 8, 672, 2002. 50. Al Dahouk, S. et al. Changing epidemiology of human brucellosis, Germany, 1962–2005. Emerg. Infect. Dis., 13, 1895, 2007. 51. Eriksen, N. et al. Brucellosis in immigrants in Denmark. Scand. J. Infect. Dis., 34, 540, 2002.

Brucella 52. White Jr. A.C. and Atmar, R.L. Infections in Hispanic immigrants. Clin. Infect. Dis., 34, 1627, 2002. 53. Memish, Z.A. and Balkhy, H.H. Brucellosis and international travel. J. Travel Med., 11, 49, 2004. 54. Troy, S.B., Rickman, L.S. and Davis, C.E. Brucellosis in San Diego. Epidemiology and species-related differences in acute clinical presentations. Medicine, 84, 174, 2005. 55. Farrell, I.D. The development of a new selective medium for the isolation of Brucella abortus from contaminated sources. Res. Vet. Sci., 16, 280, 1974. 56. Castaneda, M.R. A practical method for routine blood cultures in brucellosis. Proc. Soc. Exp. Biol. Med., 64, 114, 1947. 57. Barham, W.B. et al. Misidentification of Brucella species with use of rapid bacterial identification systems. Clin. Infect. Dis., 17, 1068, 1993. 58. Al Dahouk, S. et al. Laboratory-based diagnosis of brucellosis – a review of the literature. Part II: Serological tests for brucellosis. Clin. Lab., 49, 577, 2003. 59. OIE (World Organisation for Animal Health). Bovine brucellosis. In OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals, 5th ed. OIE, Paris, 2004. 60. Gall, D. and Nielsen, K. Serological diagnosis of bovine brucellosis: A review of test performance and cost comparison. Rev. Sci. Technol., 23, 989, 2004. 61. OIE (World Organisation for Animal Health). Caprine and ovine brucellosis (excluding Brucella ovis). In OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals, 5th ed. OIE, Paris, 2004. 62. Godfroid, J. et al. How to substantiate eradication of bovine brucellosis when aspecific serological reactions occur in the course of brucellosis testing. Vet. Microbiol., 90, 461, 2002. 63. Nielsen, K. Diagnosis of brucellosis by serology. Vet. Microbiol., 90, 447, 2002. 64. Pouillot, R. et al. The brucellin skin test as a tool to differentiate false positive serological reactions in bovine brucellosis. Vet. Res., 28, 365, 1997. 65. Saergerman, C. et al. Diagnosis of bovine brucellosis by skin test: conditions for the test and evaluation of its performance. Vet. Rec., 145, 214, 1999. 66. Morgan, W.J.B. et al. The rose bengal plate agglutination test in the diagnosis of brucellosis. Vet. Rec., 85, 636, 1969. 67. Patterson, J.M., Deyoe, B.L. and Stone, S.S. Identification of immunoglobulins associated with complement fixation, agglutination, and low pH buffered antigen tests for brucellosis. Am. J. Vet. Res., 37, 319, 1976. 68. Rose, J.E. and Roepke, M.H. An acidified antigen for detection of nonspecific reactions in the plate-agglutination test for bovine brucellosis. Am. J. Vet. Res., 18, 550, 1957. 69. Alton, G.G. et al. The serological diagnosis of bovine brucellosis: an evaluation of the complement fixation, serum agglutination and Rose Bengal tests. Aust. Vet. J., 51, 57, 1975. 70. Stemshorn, B.W. et al. A comparison of standard serological tests for the diagnosis of bovine brucellosis in Canada. Can. J. Comp. Med., 49, 391, 1985. 71. Jones, L.M., Hendricks, J.B. and Berman, D.T. The standardization and use of the complement fixation test for the diagnosis of bovine brucellosis, with a review of the literature. Am. J. Vet. Res., 24, 1143, 1963. 72. Nicoletti, P. and Muraschi, T.F. Bacteriologic evaluation of serologic test procedures for the diagnosis of brucellosis in problem cattle herds. Am. J. Vet. Res., 27, 689, 1966. 73. Nielsen, K. et al. Improved competitive enzyme immunoassay for the diagnosis of bovine brucellosis. Vet. Immunol. Immunopathol., 46, 285, 1995.

329 74. Nielsen, K.H. et al. Comparison of enzyme immunoassays for the diagnosis of bovine brucellosis. Prev. Vet. Med., 26, 17, 1996. 75. Nielsen, K. et al. A homogeneous fluorescence polarisation assay for detection of antibody to Brucella abortus. J. Immunol. Methods, 195, 161, 1996. 76. Samartino, L. et al. Fluorescence polarization assay: Appli­ cation to the diagnosis of bovine brucellosis in Argentina. J. Immunoassay, 20, 115, 1999. 77. Nielsen, K. et al. Fluorescence polarization assay for the diagnosis of bovine brucellosis: adaptation to field use. Vet. Microbiol., 80, 163, 2001. 78. Bricker, B.J. PCR as a diagnostic tool for brucellosis. Vet. Microbiol., 90, 435, 2002. 79. Leal-Klevezas, D.S. et al. Single-step PCR for detection of Brucella spp. from blood and milk of infected animals. J. Clin. Microbiol., 12, 3087, 1995. 80. O’Leary, S., Sheahan, M. and Sweeney, T. Brucella abortus detection by PCR assay in blood, milk and lymph tissue of serologically positive cows. Res. Vet. Sci., 81, 170, 2006. 81. Romero, C. and López-Goni, I. Improved method for purification of bacterial DNA from bovine milk for detection of Brucella spp. by PCR. Appl. Environ. Microbiol., 65, 3735, 1999. 82. Romero, C. et al. Evaluation of PCR and indirect enzymelinked immunosorbent assay on milk samples for diagnosis of brucellosis in dairy cattle. J. Clin. Microbiol., 33, 3198, 1995. 83. Gallien, P. et al. Detection of Brucella species in organs of naturally infected cattle by polymerase chain reaction. Vet. Rec., 142, 512, 1998. 84. Serpe, L. et al. Single-step method for rapid detection of Brucella spp. in soft cheese by gene-specific polymerase chain reaction. J. Dairy Res., 66, 313, 1999. 85. Tantillo, G. et al. Polymerase chain reaction for the direct detection of Brucella spp. in milk and cheese. J. Food. Prot., 64, 164, 2001. 86. Tantillo, G.M., Di Pinto, A. and Buonavoglia, C. Detection of Brucella spp. in soft cheese by semi-nested polymerase chain reaction. J. Dairy Res., 70, 245, 2003. 87. Hamdy, M.E.R. and Amin, A.S. Detection of Brucella species in the milk of infected cattle, sheep, goats and camels by PCR. Vet. J., 163, 299, 2002. 88. Leal-Klevezas, D.S. et al. Use of polymerase chain reaction to detect Brucella abortus biovar 1 in infected goats. Vet. Microbiol., 75, 91, 2000. 89. Herman, L. and De Ridder, H. Identification of Brucella spp. by using the polymerase chain reaction. Appl. Environ. Microbiol., 58, 2099, 1992. 90. Romero, C. et al. Specific detection of Brucella DNA by PCR. J. Clin. Microbiol., 3, 615, 1995. 91. Rijpens, N.P. et al. Direct detection of Brucella spp. in raw milk by PCR and reverse hybridization with 16S-23S rRNA spacer probes. Appl. Environ. Microbiol., 62, 1683, 1996. 92. Leal-Klevezas, D.S., Lopéz-Merino, A. and Martínez-Soriano, J.P. Molecular detection of Brucella spp.: Rapid identification of B. abortus biovar 1 using PCR. Arch. Med. Res., 26, 263, 1995. 93. Ouahrani, S. et al. Identification and sequence analysis of IS6501, an insertion sequence in Brucella spp.: Relationship between genomic structure and the number of IS6501 copies. J. Gen. Microbiol., 139, 3265, 1993. 94. Ouahrani-Bettache, S., Soubrier, M.P. and Liautard, J.P. IS6501-anchored PCR for detection and identification of Brucella species and strains. J. Appl. Bacteriol., 81, 154, 1996.

330 95. Baily, G.G. et al. Detection of Brucella melitensis and Brucella abortus by DNA amplification. J. Trop. Med. Hyg., 95, 271, 1992. 96. Bricker, B.J. and Halling, S.M. Differentiation of Brucella abortus bv. 1, 2, and 4, Brucella melitensis, Brucella ovis, and Brucella suis bv. 1 by PCR. J. Clin. Microbiol., 11, 2660, 1994. 97. García-Yoldi, D. et al. Multiplex PCR assay for the identification of all Brucella species and the vaccine strains Brucella abortus S19 and RB51 and Brucella melitensis Rev1. Clin. Chem., 52, 779, 2006. 98. Al Dahouk, S. et al. The detection of Brucella spp. using PCR-ELISA and real-time PCR assays. Clin. Lab., 50, 387, 2004. 99. Al Dahouk, S. et al. Evaluation of genus-specific and speciesspecific real-time PCR assays for the identification of Brucella spp. Clin. Chem. Lab. Med., 45, 1464, 2007. 100. Redkar, R. et al. Real-time detection of Brucella abortus, Brucella melitensis and Brucella suis. Mol. Cell. Probes, 15, 43, 2001. 101. Newby, D.T., Hadfield, T.L. and Roberto, F.F. Real-time PCR detection of Brucella abortus: A comparative study of SYBR green I, 5´-exonuclease and hybridization probe assays. Appl. Environ. Microbiol., 69, 4753, 2003. 102. Probert, W.S. et al. Real-time multiplex PCR assay for detection of Brucella spp., B. abortus, and B. melitensis. J. Clin. Microbiol., 42, 1290, 2004. 103. Foster, J.T. et al. Real-time PCR assays of single-nucleotide polymorphisms defining the major Brucella clades. J. Clin. Microbiol., 46, 296, 2008. 104. Mukherjee, F. et al. Multiple genus-specific markers in PCR assays improve the specificity and sensitivity of diagnosis of brucellosis in field animals. J. Med. Microbiol., 56, 1309, 2007. 105. van Belkum, A. High-throughput epidemiologic typing in clinical microbiology. Clin. Microbiol. Infect., 9, 86, 2003. 106. Gee, J.E. et al. Use of 16S rRNA gene sequencing for rapid confirmatory identification of Brucella isolates. J. Clin. Microbiol., 42, 3649, 2004. 107. Halling, S.M. et al. Completion of the genome sequence of Brucella abortus and comparison to the highly similar genomes of Brucella melitensis and Brucella suis. J. Bacteriol., 187, 2715, 2005. 108. Fekete, A. et al. Amplification fragment length polymorphism in Brucella strains by use of polymerase chain reaction with arbitrary primers. J. Bacteriol., 23, 7778, 1992. 109. Mercier, E. et al. Polymorphism in Brucella strains detected by studying distribution of two short repetitive DNA elements. J. Clin. Microbiol., 5, 1299, 1996. 110. Tscherneva, E. et al. Repetitive element sequence based polymerase chain reaction for typing of Brucella strains. Vet. Microbiol., 51, 169, 1996. 111. Tscherneva, E. et al. Differentiation of Brucella species by random amplified polymorphic DNA analysis. J. Appl. Microbiol., 88, 69, 2000. 112. Al Dahouk, S. et al. Identification of Brucella species and biotypes using polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP). Crit. Rev. Microbiol., 31, 191, 2005. 113. Whatmore, A.M. et al. Use of amplified fragment length polymorphism to identify and type Brucella isolates of medical and veterinary interest. J. Clin. Microbiol., 43, 761, 2005.

Molecular Detection of Foodborne Pathogens 114. Marianelli, C. et al. Molecular characterization of the rpoB gene in Brucella species: New potential molecular markers for genotyping. Microbes Infect., 8, 860, 2006. 115. Scott, J.C. et al. Multiplex assay based on single-nucleotide polymorphisms for rapid identification of Brucella isolates at the species level. Appl. Environ. Microbiol., 73, 7331, 2007. 116. Whatmore, A.M., Perrett, L.L. and MacMillan, A.P. Characterisation of the genetic diversity of Brucella by multilocus sequencing. BMC Microbiol., 7, 34, 2007. 117. DelVecchio, V.G. et al. The genome sequence of the facultative intracellular pathogen Brucella melitensis. Proc. Natl. Acad. Sci. USA, 99, 443, 2002. 118. Paulsen, I.T. et al. The Brucella suis genome reveals fundamental similarities between animal and plant pathogens and symbionts. Proc. Natl. Acad. Sci. USA, 99, 13148, 2002. 119. Al Dahouk, S. et al. Evaluation of Brucella MLVA typing for human brucellosis. J. Microbiol. Methods, 69, 137, 2007. 120. Le Flèche, P. et al. Evaluation and selection of tandem repeat loci for a Brucella MLVA typing assay. BMC Microbiol., 6, 9, 2006. 121. Whatmore, A.M. et al. Identification and characterization of variable-number of tandem-repeat markers for typing of Brucella spp. J. Clin. Microbiol., 44, 1982, 2006. 122. Vergnaud, G. and Denoeud, F. Minisatellites: Mutability and genome architecture. Genome Res., 10, 899, 2000. 123. Bricker, B.J. and Ewalt, D.R. Evaluation of the HOOF-print assay for typing Brucella abortus strains isolated from cattle in the United States: Results with four performance criteria. BMC Microbiol., 5, 37, 2005. 124. Bricker, B.J., Ewalt, D.R. and Halling, S.M. Brucella ‘HOOF-prints’: Strain typing by multi-locus analysis of variable number tandem repeats (VNTRs). BMC Microbiol., 3, 15, 2003. 125. Casanas, M.C. et al. Specificity of a polymerase chain reaction assay of a target sequence on the 31-kilodalton Brucella antigen DNA used to diagnose human brucellosis. Eur. J. Clin. Microbiol. Infect. Dis., 20, 127, 2001. 126. Kattar, M.M. et al. Development and evaluation of real-time polymerase chain reaction assays on whole blood and paraffin-embedded tissues for rapid diagnosis of human brucellosis. Diagn. Microbiol. Infect. Dis., 59, 23, 2007. 127. Queipo-Ortuno, M.I. et al. Comparison of seven commercial DNA extraction kits for the recovery of Brucella DNA from spiked human serum samples using real-time PCR. Eur. J. Clin. Microbiol. Infect. Dis., 27, 109, 2008. 128. Navarro, E. et al. Use of real-time quantitative polymerase chain reaction to monitor the evolution of Brucella melitensis DNA load during therapy and post-therapy follow-up in patients with brucellosis. Clin. Infect. Dis., 42, 1266, 2006. 129. Christensen, D.R. et al. Detection of biological threat agents by real-time PCR: Comparison of assay performance on the R.A.P.I.D., the LightCycler, and the Smart Cycler platforms. Clin. Chem., 52, 141, 2006. 130. Debeaumont, C., Falconnet, P.A., and Maurin, M. Realtime PCR for detection of Brucella spp. DNA in human serum samples. Eur. J. Clin. Microbiol. Infect. Dis., 24, 842, 2005. 131. Queipo-Ortuno, M.I. et al. Rapid diagnosis of human brucellosis by SYBR green I-based real-time PCR assay and melting curve analysis in serum samples. Clin. Microbiol. Infect., 11, 713, 2005.

24 Burkholderia

Karlene H. Lynch and Jonathan J. Dennis University of Alberta

Contents 24.1 Introduction.....................................................................................................................................................................331 24.1.1 Burkholderia pseudomallei, Burkholderia mallei, and the BCC......................................................................331 24.1.1.1 Burkholderia pseudomallei and Burkholderia mallei......................................................................331 24.1.1.2 BCC.................................................................................................................................................. 332 24.1.2 Burkholderia gladioli pathovar cocovenenans................................................................................................. 333 24.1.2.1 Taxonomy......................................................................................................................................... 333 24.1.2.2 Epidemiology................................................................................................................................... 333 24.1.2.3 Pathogenesis..................................................................................................................................... 334 24.1.2.4 Biology............................................................................................................................................. 335 24.1.2.5 Conventional Diagnosis................................................................................................................... 336 24.1.2.6 Molecular Diagnosis........................................................................................................................ 337 24.2 Methods.......................................................................................................................................................................... 337 24.2.1 Reagents and Equipment................................................................................................................................... 337 24.2.2 Burkholderia gladioli pathovar cocovenenans Sample Collection and Preparation........................................ 338 24.2.2.1 Bacterial Isolation, Propagation, and Storage.................................................................................. 338 24.2.2.2 Toxin Extraction............................................................................................................................... 338 24.2.3 Burkholderia gladioli pathovar cocovenenans Detection Procedure............................................................... 338 24.3 Conclusions and Future Perspectives............................................................................................................................. 339 References.................................................................................................................................................................................. 340

24.1  Introduction The genus Burkholderia is an incredibly diverse group of Gram-negative β-proteobacteria. Although there are currently at least 60 species and proposed species in this genus, very few of these have been studied extensively.1 Much of the research to date has focused on the Burkholderia cepacia complex (BCC), Burkholderia pseudomallei, Burkholderia mallei, and Burkholderia gladioli. BCC organisms are pathogens that cause serious infections in plants, animals, and humans.2–4 However, they can also be beneficial as they fix nitrogen, produce antibiotics and antifungals, and degrade organic compounds.5–7 B. pseudomallei causes melioidosis, a disease with a wide variety of symptoms, while B. mallei causes glanders, a condition usually found in horses that can also affect humans.8 Since 2003, B. gladioli has been divided into four pathovars: gladioli, alliicola, agaricicola, and cocovenenans.9 The first three groups are primarily plant pathogens.10,11 However, in addition to being phytopathogenic, members of these pathovars can also infect patients with chronic granulomatous disease (CGD), cystic fibrosis (CF), and acquired immune deficiency syndrome (AIDS).9,12,13 The taxonomic description of B. gladioli pvs. gladioli, alliicola, and agaricicola published in 2003 indicates that they do not produce toxins that are harmful to humans, although some strains have since been shown to synthesize toxoflavin (discussed below).9,14

The fourth pathovar, B. gladioli pv. cocovenenans, is genetically similar to the first three pathovars, but distinct with regards to both its epidemiology and pathogenicity. Although other Burkholderia species can be found in food and water supplies (including B. pseudomallei, B. mallei, and the BCC), B. gladioli pv. cocovenenans is the only division of the Burkholderia genus that is characterized as a foodborne pathogen. In an analogous fashion to Clostridium botulinum, these bacteria cause disease indirectly through the production of toxins that permeate contaminated foods. This chapter will briefly discuss the literature regarding the presence of B. pseudomallei, B. mallei, and the BCC in food and water supplies and the potential for transmission from these sources, followed by an extensive review of B. gladioli pv. cocovenenans epidemiology, pathogenesis, biology, and diagnostics.

24.1.1 Burkholderia pseudomallei, Burkholderia mallei, and the BCC 24.1.1.1 Burkholderia pseudomallei and Burkholderia mallei B. pseudomallei causes melioidosis, a potentially fatal condition with a variety of symptoms including pneumonia, skin lesions, and septic shock.8 Most melioidosis cases are seen in 331

332

Southeast Asia (especially Thailand) and northern Australia, although they can also be found in areas such as Brazil.8,15 Although the first cases of melioidosis were described in 1913, there remains some debate as to how B. pseudomallei is transmitted to humans.16,17 It is thought that most infections occur following either inoculation of broken skin or inhalation.18 Historically, it was considered that melioidosis was also spread through the consumption of contaminated food and water, as it could be transmitted in this manner to guinea pigs, rabbits, and rats.16,19 However, B. pseudomallei has not been definitively shown to spread to humans by this route. A report published in 1998 described the development of melioidosis in five adults in northwestern Australia.20 Using pulsed-field gel electrophoresis (PFGE) typing, it was determined that all five patients were infected with the same PFGE type and that this type was identical to that of B. pseudomallei isolates found in the area’s drinking water. Although this association suggests that these cases developed due to ingestion of the waterborne microbe, the possibility exists that these persons may have been infected via a different route.20 B. pseudomallei can also be isolated from drinking water in areas where melioidosis is not endemic. Zanetti et al. showed that B. pseudomallei could be isolated from 7.1% of drinking water samples taken in Bologna, Italy.21 In these samples, the levels of B. pseudomallei ranged from 1020 to 15000 colony forming units (CFU) per 100 ml sample. Treatment of water with chlorine will kill B. pseudomallei (as well as B. mallei) in an experimental setting, but various factors in the environment (including attachment and nutrient limitation) may affect this susceptibility.22 B. mallei causes glanders, a disease of horses that can also be transmitted to humans. The symptoms vary ­depending on the route of transmission, but can include pneumonia, skin lesions, and septic shock (similar to melioidosis).23 Horses generally become infected following ingestion of B. ­mallei introduced into their food and water by other infected ­horses.23 It is thought that humans may also become infected via ingestion.24 As a result of stringent measures introduced to control the spread of glanders, this condition has not been seen in Western countries since 1939 (except following accidental laboratory exposure).25,26 However, there are concerns that both B. pseudomallei and B. mallei could be released as bioterrorism agents.27 Both of these species have been identified as food defense concerns.28 This categorization indicates that these species, both of which are Category B biological agents based on Centers for Disease Control classification, could be purposely introduced into the food supply in an act of terrorism.27,28 This is in contrast with normal concerns over food safety in which organisms such as Streptococcus may lead to accidental contamination of the food supply.28 24.1.1.2  BCC The BCC is currently made up of 15 (atleast 17) closely related Burkholderia species.1 These bacteria cause potentially fatal infections in susceptible patients, particularly those with CF and CGD.29,30 Whereas both melioidosis and glanders can have a wide range of symptoms that affect many different

Molecular Detection of Foodborne Pathogens

systems of the body, BCC infections tend to be ­limited to the lungs. These infections may be asymptomatic, may cause a patient’s condition to worsen over time, or may develop into a rapidly fatal pneumonia accompanied by damage to the lung tissue and septicemia.4 This group of bacteria is found more commonly in food and water supplies than either B. pseudomallei or B. mallei. They can be isolated from the rhizospheres of rice, corn, wheat, soybean, and alfalfa plants.31,32 BCC bacteria have also been found associated with Pleurotus ostreatus, the oyster mushroom, which is the second-most cultivated mushroom in the United States.33,34 BCC organisms may cause spoilage in foods as diverse as onions, Swiss cheese, and Parma ham.2,35,36 However, these bacteria can also prevent food spoilage: a BCC strain isolated from Washington navel oranges was found to prevent fungal infection of these fruits during storage when they were treated with this strain.37 Because of the ability of BCC organisms to produce such antifungals and antibiotics (in addition to their ability to fix nitrogen), there has been a great deal of interest in developing these species into agricultural products. However, because of the potential risk to susceptible persons, the United States Environmental Protection Agency (EPA) has placed strict limitations on the production and use of BCC bacteria since 2003.38 Despite their wide distribution in food products, when Moore et al. assessed the prevalence of BCC bacteria in a variety of foods (including bakery items, onions, cheese, ham, and oranges), they were only able to isolate these organisms from unpasteurized milk.39 BCC species have also been isolated from unpasteurized milk by Uraz and Çitak and Munsch-Alatossava and Alatossava.40–43 It is unknown how the milk becomes contaminated, but it has been suggested that it is due to transfer of BCC organisms from the soil to the cow udder and/or to milk storage tanks.44 Berriatua et al. found that the BCC species Burkholderia cenocepacia and Burkholderia vietnamiensis were responsible for causing subclinical mastitis (inflammation of the udder) in milking sheep.3 However, when Moore et al. examined milk samples taken from cows with mastitis to determine if this condition was responsible for the contamination, they were unable to isolate BCC organisms.44 It is important to note that, despite the contamination of raw milk, these bacteria are not found in commercially available dairy products and are effectively killed by pasteurization.39,44 Like B. pseudomallei, BCC organisms can also be isolated from drinking water. Zanetti et al. found that 3.5% of drinking water samples in Bologna, Italy contained BCC bacteria.21 The counts in these samples were very low compared to B. pseudomallei, containing between 1 and 19 CFU per 100 ml sample. Pegues et al. examined a BCC isolate from a water jug at a CF summer camp.45 They found that the ribotype of this isolate was different from the ribotypes taken from patients infected at the camp, indicating that this water was not the source of infection. The presence of BCC organisms in these samples may be due to their propensity to form biofilms in water supply systems.46 In contrast to these

Burkholderia

findings, examination of samples taken either by Moore et al. or Vermis et al. were unable to detect BCC organisms in a variety of drinking water sources, including bottled, tap, well, and spring water.39,47 Although it is generally considered that BCC organisms do not spread through ingestion, it remains a possibility that if these bacteria are present in the mouth and throat following the consumption of contaminated food or water, they may subsequently enter the lungs.39 Therefore, it is recommended that patients with CF should not consume unpasteurized milk products or food from buffets and should thoroughly wash and dry produce.39,48,49 To date, only one study has suggested a link between food consumption and BCC infection in CF patients. Fisher et al. found that BCC isolates taken from cucumber, lettuce, and cauliflower purchased from salad bars and food stores near two CF centers were of the same ribotype that was most commonly isolated from the patients in these centers.50 However, it is not possible to discern from these findings whether or not these foods acted as sources of infection.

24.1.2 Burkholderia gladioli pathovar cocovenenans

24.1.2.1  Taxonomy The taxonomy of the bacteria currently named B. gladioli pv. cocovenenans is relatively complicated and incredibly plastic. Despite the publication of several articles defining the nomenclature of this group, the naming remains inconsistent in the current literature. van Veen and Mertens published the first studies on this bacterium, which they isolated from contaminated tempe bongkrek, an Indonesian fermented coconut cake.51 This organism was named Pseudomonas cocovenenans (cocos=coconut, veneno=to poison) and this nomenclature was retained in early studies.52–54 Around this time, another bacterium was isolated from contaminated fermented corn flour in China.55 Similar isolates were studied extensively in the Chinese literature, but were not described in an English journal until 1988. Here, the organism is identified as Flavobacterium farinofermentans sp. nov.56 However, in the Chinese literature, these bacteria had been called by this name since 1980, and had already been renamed Pseudomonas farinofermentans by the mid-1980s.57,58 Comparisons of F. farinofermentans sp. nov. and P. cocovenenans (published in Chinese in 1987 and cited in the English literature in 1989) suggested that F. farinofermentans sp. nov. be renamed P. cocovenenans subspecies farinofermentans.59,60 A similar conclusion was reached following a separate set of comparisons published in 1988 and 1990, first in Chinese and then in English. After assessing a variety of characteristics (including cell and colony morphology, substrate utilization, antibiotic resistance, DNA hybridization, and antibody cross-reactivity), Zhao et al. determined that P. farinofermentans and P. cocovenenans belonged to the same species.58,61 However, they suggested that, because there were slight differences observed between P. farinofermentans and P. cocovenenans isolates

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with respect to substrate utilization and antibiotic resistance, P.  farinofermentans should be renamed P. cocovenenans biovar farinofermentans. In 1992, seven Pseudomonas species were reclassified and assigned to the new genus Burkholderia (including B. pseudomallei, B. mallei, B. cepacia, and B. gladioli).62 Following analysis of the type strain LMG 11626/NCIB 9450/ATCC 33664 (a strain isolated from tempe bongkrek), P. cocovenenans was subsequently assigned to this genus in 1995, becoming Burkholderia cocovenenans comb. nov.63 This transfer was confirmed later that year using 16S rDNA sequencing using NCIMB 12451 (a fermented corn flour poisoning isolate).64 In this paper, this strain was described as P. cocovenenans (as opposed to P. cocovenenans bv. farinofermentans), thus discontinuing the use of the biovar designation. In 1997, Vandamme et al. found that, when SDS-PAGE of whole-cell proteins was performed using B. gladioli and B. cocovenenans strains, the resulting protein patterns were very similar.65 To confirm the relatedness of these two species, this group performed further experiments using SDSPAGE, DNA hybridization, and biochemical tests.66 Their results led to the reclassification of B. cocovenenans as a member of B. gladioli. The most recent change to this taxonomy has been the addition of a fourth pathovar to B. gladioli that encompasses the former B. cocovenenans strains and isolates, called B. gladioli pathovar cocovenenans. The three existing pathovars of B. gladioli (gladioli, alliicola, and agaricicola) are all phytopathogenic.10,11 These pathovars can be differentiated from pathovar cocovenenans because they do not produce bongkrekic acid (discussed below) and have slightly different substrate utilization patterns (Table 24.1).9 Despite the publication of the papers described here, discrepancies in taxonomy remain in the current literature, particularly with regards to Chinese publications where use of the name P. cocovenenans subsp. farinofermentans remains common.67–70 24.1.2.2  Epidemiology B. gladioli pv. cocovenenans intoxications occur in two areas of the world due to the consumption of two different locally-produced fermented foods. The first of these foods, from which the bacterium was initially isolated, is tempe (or tempeh) bongkrek (also referred to as semaji or bongkrek). Tempe refers to a group of foods made by fermentation of plant material (such as soybeans and coconut) using mould.71 Tempe bongkrek is made using coconut presscake (a byproduct of processing coconut to remove the oil) or coconut milk residue.71,72 This material is then fermented using Rhizopus oligosporus, a fungus that grows on the surface of and throughout the tempe.73 The finished product is a white cake with a thickness of one inch or less.74 Tempe bongkrek that is properly made and fully fermented tends to be very safe.74 However, when it is made in the home (often by those who have little knowledge or experience regarding proper production methods), insufficient fungal growth may allow bacteria—including B. gladioli pv. cocovenenans—to multiply.74

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Molecular Detection of Foodborne Pathogens

Table 24.1 Differential Characteristics Among Pathovars of B. gladioli B. gladioli Pathovar gladioli NCPPB 1891

Pathovar agaricicola NCPPB 3580

Pathovar alliicola NCPPB 947

Pathovar cocovenenans ATCC 33664 and 6 Strains

Gladiolus – +

Mushroom – –

Onion – +

Coconut, fermented corn 100% (7/7)* 100% (7/7)

Growth at 4°C Nitrate reduction

+

+

–

0% (0/7)

–

–

+

100% (7/7)

Xylitol Gentiobiose N-Acetyl-D-galactosamine Acetic acid L-Pyroglutamic acid Maltose D-Raffinose Sucrose Glycyl-L-aspartic acid

– + + + – – – – –

Assimilation – + + – + + – – –

– + + + + – + + +

100% (7/7) 0% (0/7) 0% (0/7) 86% (6/7) 100% (7/7) 0% (0/7) 0% (0/7) 0% (0/7) 0% (0/7)

Characteristics Source Bongkrekic acid production Growth at 41°C

* % of positive strains; +: positive or weak positive reaction, –: negative reaction. Source: Jiao, Z. et al., Microbiol. Immunol., 47, 915, 2003. Reproduced with permission from Wiley-Blackwell Publishing.

Most tempe bongkrek poisonings occur in the Banjumas province in Central Java, Indonesia.74 Here, tempe is an important food source because it is inexpensive and highly nutritious, containing both protein and vitamin B12.52,71 The first reports of tempe bongkrek poisoning date back to 1895.75 Since then, poisonings have been relatively frequent, with 7216 cases and 850 deaths occurring between 1951 and 1975.74,76 Despite the banning of tempe bongkrek by the government in 1988 (and the threat of prison time), Indonesians continue to make and consume this food.71,77 Although there have been no reports of intoxications in surrounding countries, this is not necessarily an indication that they have not occurred.72 The second product that can cause B. gladioli pv. cocovenenans intoxications is fermented corn flour (FCF). FCF is produced by grinding corn that has been immersed in water for between 2 and 4 weeks. This flour is then used to make other foods, including breads and noodles.78 B. gladioli pv. cocovenenans can grow in the flour during storage.73 Cooking or baking of contaminated foods does not prevent illness as the toxins produced by the bacteria are heat-stable.74 FCF is commonly made in northeastern China, including the provinces of Heilongjiang, Jilin, and Liaoning.78 Intoxications due to B. gladioli pv. cocovenenans occur relatively frequently, with 226 outbreaks identified in these provinces between 1953 and 1975. Of the 1842 patients affected, 703 died (38.2% mortality). Most intoxications occurred in the summer months, between July and September. There was no effect of age or sex on whether or not those who ate contaminated FCF became ill or died. These effects were instead related to the amount of toxic material that had been eaten.78 A second,

less common source of B. gladioli pv. cocovenenans intoxications in China (and other Asian countries) is Tremella fuciformis, an edible mushroom. It has been found that up to half of these mushrooms may be contaminated with B. gladioli pv. cocovenenans.64 Following consumption of food contaminated with B. gladioli pv. cocovenenans, intoxication occurs rapidly, usually within 1–10 h.78 The classical symptoms are hyperglycemia followed by hypoglycemia, spasms, loss of consciousness, and death.74 Following autopsy, the liver, kidneys, brain, heart, lungs, gastrointestinal tract, and spleen all show signs of damage, particularly the first three organs.78 When food contaminated with B. gladioli pv. cocovenenans is fed to dogs, they develop spasms, enter comas and die within 2–3 h.78 Dogs and rhesus monkeys fed B. gladioli pv. cocovenenans culture supernatants die within 6–33 h and 15.5–35 h, respectively.56 When mice are fed the B. gladioli pv. cocovenenans toxin bongkrekic acid, all animals die within 45 minutes.9 24.1.2.3  Pathogenesis Although B. pseudomallei, B. mallei, and BCC species express a wide variety of virulence factors (including exopolysaccharide, type III secretion systems, and lipopolysaccharide), no single factor has been shown to have a predominant effect.8,23,79 This is in contrast to B. gladioli pv. cocovenenans, an organism whose pathogenicity is mediated solely by toxins. When mice are administered either live or heat-killed bacteria, adverse effects are not observed, as opposed to when they are administered culture supernatants or partiallypurified toxin.56,9 B. gladioli pv. cocovenenans produces two

Burkholderia

toxins, toxoflavin (also called xanthothricin) and bongkrekic acid (also called bongkrek acid or flavotoxin A). Toxoflavin has the chemical formula C7H7N5O2. This toxin is not restricted to B. gladioli pv. cocovenenans, as it is also produced by Streptomyces, Burkholderia glumae, Burkholderia plantarii, and phytopathogenic B. gladioli.80–82,14 In mice, the LD50 for injected toxoflavin is 1.7 mg/kg, while for the ingested toxin it is 8.4 mg/kg.83 This compound has a bright yellow color, a melting point of approximately 170°C, and a UV absorption maximum of 258 nm.54 Tempe bongkrek with a yellow color is particularly dangerous as it has been permeated by this toxin and likely by bongkrekic acid as well.72,84 However, both toxins may still be present even if there is no change in coloration.84 Several papers have been published describing the chemical synthesis of toxoflavin.85–87 This substance is toxic because it acts as an electron carrier, resulting in the production of hydrogen peroxide. Under normal conditions, NADH, and FADH2 produced during glycolysis and the citric acid cycle (in the cytosol and mitochondrial matrix, respectively) transfer their electrons to a series of carriers in the inner mitochondrial membrane, including cytochromes, which are organized into Complex I–IV.88 The electrons are shuttled down this chain based on redox potential, and the energy released is used to pump protons into the intermembrane space of the mitochondria. This transport creates a proton gradient that powers the ATP synthase, which produces the majority of ATP used by the cell in a process called oxidative phosphorylation.88 It was discovered by Latuasan and Berends that toxoflavin can interfere with this process.89 When antimycin A, which blocks Complex III of the electron transport chain, is added to yeast, respiration is inhibited.88,89 However, when antimycin A and toxoflavin are added together, no inhibitory effects are observed. Similarly, the addition of potassium cyanide, which blocks Complex IV, inhibited respiration, but not in the presence of toxoflavin.88,89 These results suggest that toxoflavin allows for cytochrome-independent electron transfer.89 Toxoflavin also exhibits electron-transferring activity independent of the mitochondrion. Latuasan and Berends showed that this toxin was able to stimulate oxygen uptake by both a mitochondria-free yeast cell extract and a yeast strain that lacks cytochrome proteins.89 When NADH is combined with toxoflavin in a yeast cell extract, electrons are transferred from NADH to toxoflavin and toxoflavin is reduced. Toxoflavin then transfers these electrons to oxygen, producing hydrogen peroxide. It is predicted that the toxic effect of toxoflavin is due to the action of the hydrogen peroxide and not due to interference with the electron transport chain.89 Yeast cells (which exhibit very high catalase activity) do not show growth defects in the presence of toxoflavin (even though their electron transport system is altered), which would be expected if interference with the mitochondria were contributing to toxicity.89 Although toxoflavin is highly toxic, it is not generally the key agent in B. gladioli pv. cocovenenans intoxications. Instead, the toxin with the greatest clinical relevance is

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bongkrekic acid (especially in the case of tempe bongkrek), as it is both more toxic and present in higher concentrations in contaminated foods.83,84 The LD50 in mice is 1.4 mg/kg when injected and 3.16 mg/kg when ingested.83,60 This compound has the chemical formula C28H38O7. Several papers have been published that have determined the properties and structure of bongkrekic acid.83,90–92 This compound is toxic because it inhibits oxidative phosphorylation by binding to the adenine nucleotide transporter (responsible for the shuttling of ADP and ATP across the inner mitochondrial membrane). The mechanism of action of bongkrekic acid was first examined in 1960. While the addition of toxoflavin to a cell homogenate sharply increases oxygen consumption, addition of bongkrekic acid has the opposite effect.89,93 Tests performed on isolated rat heart mitochondria by Welling et al. indicated that the oxidation of pyruvate (the end product of glycolysis), α-ketoglutarate, and malate (two citric acid cycle intermediates) were inhibited by bongkrekic acid.93 The mechanism behind this inhibition was not elucidated until 1970. Henderson and Lardy found that, when rat liver mitochondria were treated with bongkrekic acid, these organelles were no longer able to take up 14C-labeled ADP or ATP, suggesting that bongkrekic acid blocks the mitochondrial adenine nucleotide transporter.94 Further experiments indicated that bongkrekic acid binds to the transporter when it is in the matrix or m-state (the conformation in which the ADP/ATP binding site faces inwards towards the matrix).95 Because bongkrekic acid inhibits oxidative phosphorylation, the only way for cells to generate ATP is through anaerobic glycolysis.96 This metabolic change is responsible for the symptoms of intoxication. Glycogen stores are broken down to allow for an increased level of glycolysis, leading to hyperglycemia and increased lactic acid levels, but these stores are soon depleted, resulting in hypoglycemia and an inability to regenerate ATP.93 It is the ATP depletion that is fatal and not the hypoglycemia, as the injection of glucose is insufficient to prevent death.93 24.1.2.4  Biology B. gladioli pv. cocovenenans cells are motile, aerobic, nonspore­-forming, nonencapsulated Gram-negative rods. These bacteria form smooth, round colonies (on potato dextrose agar [PDA]) that are yellow pigmented due to the production of toxoflavin. Growth can occur between 6 and 41°C, but is optimal between 30 and 37°C.61,97 Most toxin production occurs between 22 and 30°C.97 Both catalase and oxidase tests are positive, but the oxidase reaction is extremely weak. Other characteristics of these bacteria (along with other B. gladioli pathovars) are summarized in Table 24.1. Two important aspects of the biology of B. gladioli pv. cocovenenans that affect its viability and toxin production in food are its susceptibility to pH and salt concentration. These bacteria cannot grow at a pH of 4.5 or in a salt concentration of 6%.61 As such, measures to decrease the pH or increase the salt content of foods such as tempe bongkrek have been suggested as a means to make them safer. However, some of these methods have been met with resistance. For example,

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addition of the acidic leaves of Oxalis spp. (a flowering plant) to the raw material for making tempe bongkrek decreases the pH to approximately 5.5, inhibiting bacterial growth. However, this addition changes the color of the cake, which has made this preventive measure relatively unpopular.84 Buckle and Kartadarma found that the production of both toxins was inhibited to different degrees following alteration of the pH and salt content by the addition of acetic acid and sodium chloride.84 After 48 h incubation in coconut culture medium (CCM, discussed below) at a pH of 4.5 or less, a salt concentration of 2% or greater, or a combination of pH 5.5 and 1% salt, toxoflavin was not produced. However, only the combination of decreased pH and higher salt concentration (5.5 and 1%, respectively) was sufficient to inhibit the production of bongkrekic acid in this medium. In laboratorysynthesized tempe bongkrek, a pH of 4.5 and 2% salt concentration was sufficient to inhibit the production of both toxoflavin and bongkrekic acid by three B. gladioli pv. cocovenenans strains. R. oligosporus growth was not inhibited under these conditions. Another important aspect of B. gladioli pv. cocovenenans biology with regards to its ability to produce toxin is its response to lipid content. Garcia et al. found that B. gladioli pv. cocovenenans could grow on rich coconut media (RCM) from which lipids had been extracted, but it did not produce bongkrekic acid until coconut fat was added to a level of 20%.72 When individual fatty acids were added to the media, bongkrekic acid was only produced in the presence of the saturated lauric acid (12:0), myristic acid (14:0), and palmitic acid (16:0), as well as the unsaturated oleic acid (18:1), linoleic acid (18:2), and linolenic acid (18:3). The three saturated fatty acids comprise over 70% of the total fatty acids present in coconut oil.72 Although bongkrekic acid production is affected by lipid content, toxoflavin production does not appear to be similarly affected, as it was synthesized even in the lipid-extracted RCM with no added fat.72 Therefore, intoxications that result from consuming contaminated foods with a lower fat content (including FCF and T. fuciformis) may be due to the action of toxoflavin more so than bongkrekic acid.72 However, it has also been suggested that, because corn oil has high oleic acid content, FCF may support the production of bongkrekic acid despite its relatively low lipid content.72 24.1.2.5  Conventional Diagnosis Because these intoxications have been limited to resource-poor areas and are relatively uncommon in comparison with many other foodborne pathogens, few detection measures for B. gladioli pv. cocovenenans have been described. Conventionally, these bacteria are isolated on CCM or a variant thereof. The following protocol, described by Ko et al., produces a medium that, when inoculated with R. oligosporus, approximates the properties of tempe bongkrek in a laboratory setting.98 (1) Remove 100 g meat from a fresh coconut (white layer). (2) Add 150 ml water (60–70°C).

Molecular Detection of Foodborne Pathogens

(3) Blend 2 min. Remove 200 ml liquid (using hydraulic handpress or other method). (4) Add 200 ml water (60–70°C). Repeat step 3. (5) Autoclave 60 g cakes 20 min at 110°C. In order to inoculate the cakes with mould, R. oligosporus spore suspensions (made in sterile water) are mixed with 60 g CCM cakes, poured into Petri dishes (approximately 20 g of media per dish), and incubated at 30°C for 48 h. B. gladioli pv. cocovenenans can be propagated on both CCM and laboratory-made tempe bongkrek.98 When toxoflavin is produced by bacteria growing on either of these, the media will turn a yellow color, which is a useful diagnostic criterion.84 However, the differential properties of these types of media are somewhat limited because other bacteria can produce toxoflavin and because not all B. gladioli pv. cocovenenans strains produce detectable levels of toxoflavin under laboratory conditions.14,80–82,84 However, because these intoxications have such a limited range (both with respect to the geographic area and the types of food involved), bacterial growth and toxin production on CCM or laboratory-made tempe bongkrek can be useful for propagation and preliminary analysis of B. gladioli pv. cocovenenans in food outbreaks. B. gladioli pv. cocovenenans can be identified using commercial test kits such as the Biolog GN2 system.9,42 This method, used to identify aerobic Gram-negative organisms, measures the ability of a bacterium to use a variety of carbon sources. This test can be used to differentiate pathovar cocovenenans from the other pathovars of B. gladioli.9 However, these commercial systems may give negative or conflicting results when used with these bacteria. Segonds et al. found that, using the API 20NE system, B. gladioli pv. cocovenenans could not be identified.99 Similarly, when isolates from raw milk were compared using the Biolog GN2 and API 20NE systems, one isolate was identified as B. cocovenenans by the first system and Pseudomonas fluorescens by the second.42 An effective method to detect and determine the concentration of toxins produced by B. gladioli pv. cocovenenans is high pressure liquid chromatography (HPLC). Most protocols for toxoflavin and bongkrekic acid detection are based on the methods developed by Voragen et al.100 In this protocol, toxoflavin and bongkrekic acid are extracted separately from tempe bongkrek. The UV absorbance is measured for bongkrekic acid at 267 nm and for toxoflavin at 258 nm. The extracted samples are compared to reference standards. Using this protocol, bongkrekic acid concentrations from either 2–160 µg/ml or 0.5–80 µg/ml (depending on injection volume) and toxoflavin concentrations from 1 to 300 µg/ml could be detected.100 This system effectively quantifies the levels of toxins and can detect the presence of each toxin even if they are present in a mixture. However, this method is time-consuming (as the toxins need to be extracted from the sample prior to analysis) and, more importantly, requires expensive and specialized equipment. Because these infections occur in economically-poor areas, detection methods for B. gladioli pv. cocovenenans should ideally be as simple and cost-effective as possible.

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Burkholderia

Because there is currently no selective medium for these ­bacteria, molecular techniques that can be used for rapid and effective detection during outbreaks of food intoxication are extremely important. 24.1.2.6  Molecular Diagnosis Although 16S rDNA sequencing is a useful molecular method for differentiating many bacteria, this procedure tends to be very unsuccessful for identifying members of the Burkholderia genus. For BCC species, these sequences are over 97.7% identical, which does not allow for species-level differentiation.101 Similarly, B. pseudomallei and B. mallei cannot be distinguished using this method because they have the exact same 16S rDNA sequence.102 Zhao et al. sequenced a 291 base pair segment of the 16S rDNA and found that there was a six base pair difference between B. gladioli pv. cocovenenans NCIMB 12451 and B. gladioli EY 3258.64 Viallard et al. found that the 16S sequences of the B. gladioli pv. gladioli and B. gladioli pv. cocovenenans type strains (ATCC 10248 and ATCC 33664, respectively) were 99.9% identical when a 1484 base pair region was sequenced.103 When Jiao et al. sequenced a segment with a similar length (1291 base pairs), they found that the sequence was between 99.4 and 99.7% identical between pathovar gladioli and pathovar cocovenenans strains and 98.2–99.6% identical among pathovar cocovenenans strains.9 In addition, B. gladioli pvs. gladioli and cocovenenans sequences had very high homology to those of both B. glumae and B. plantarii (98.0–99.4%). When 16S sequencing was used to identify isolates from CF patients in France, the sequence from one isolate identified as B. gladioli had only a one base pair change from the B. gladioli pv. cocovenenans reference sequence.104 These results indicate that the sequence similarity between the 16S rDNA sequences of the B. gladioli pathovars is too high to allow for adequate differentiation. As such, many studies that have identified B. gladioli in clinical specimens using this technique have not specified the pathovar to which these isolates belong.104–106 Despite this drawback, 16S sequencing has been used to putatively identify B. gladioli pv. cocovenenans in clinical isolates. In a 25-year-old patient from Thailand who produced anti-interferon γ autoantibodies, bacteria were isolated from various sites in the body, including lymphadenoid tissue and the lungs. The 16S rDNA sequences of these isolates were determined and they were identified as Burkholderia cocovenenans.107,108 To the best of our knowledge, these are the only reports in the literature of B. gladioli pv. cocovenenans causing this type of disease. Capillary electrophoresis–single-strand conformation polymorphism (CE–SSCP) analysis is a second technique that uses 16S rDNA sequence to identify B. gladioli pv. cocovenenans.109 In this method, 16S rDNA sequences are amplified using PCR primers with fluorescent labels. Capillary gel electrophoresis is performed using an automated DNA sequencer following denaturation of the PCR product. The resulting electropherograms differ based upon the conformation of the two labeled single-strand products. Using this protocol, B. gladioli pvs. cocovenenans, gladioli, and alliicola, B. glumae,

and B. plantarii all produced very similar electropherograms (which would be expected based upon the strong homology of their 16S rDNA sequences) and could be differentiated from other Gram-negative nonfermenters but not from one another. Yet another technique that uses 16S rDNA sequence to identify B. gladioli pv. cocovenenans is microarray analysis. Jin et al. developed a microarray with probes for eubacteria, for Gram-positive and Gram-negative organisms, and for specific pathogens based on their 16S sequences.110 Bacterial 16S rDNA was amplified using fluorescently-labeled primers and hybridized to the probes on the microarray. Although the microarray in this paper included a probe designed to recognize B. gladioli pv. cocovenenans, the array was not tested with this pathovar’s 16S rDNA. In a later paper, this group tested their microarray using three B. gladioli pv. cocovenenans strains and found that the microarray could successfully differentiate these strains from other foodborne pathogens.111 However, because this microarray was not tested with nonfoodborne B. gladioli, it is unknown if the other three pathovars would cross-react using this assay. Probe-based cell fishing is a technique that was developed to analyze the members of a microbial community without culturing them.112 In the first stage, biotinylated RNA probes hybridize in situ to the 23S rRNA. The cells are then incubated with paramagnetic streptavidin-coated beads. When the bacteria are run through a column in a magnetic field, cells in which the probes hybridized will stay bound, while those with no hybridized probe will not. Using the probe DIIIBcep, Stoffels et al. were able to detect the type strain of B. gladioli pv. cocovenenans.112 However, this probe was relatively nonspecific, as it also hybridized with B. cepacia, B. vietnamiensis, B. gladioli, and B. plantarii. Although several novel methods for B. gladioli detection have been developed since 1999, not all of these procedures were tested for their ability to identify food intoxication isolates.113–115 One of the protocols that was tested with B. gladioli pv. cocovenenans strains used primers specific for the gyrB gene in a multiplex PCR protocol.116 These primers amplify a 479 base pair fragment from B. gladioli pvs. cocovenenans, gladioli, alliicoli, and agaricicola, but not from B. glumae, B. plantarii, or B. cepacia. Therefore, this protocol is more specific than either CE–SSCP or probe-based cell fishing with respect to the detection of B. gladioli pv. cocovenenans. Although this method does not specifically identify this pathovar, it can still be a useful diagnostic tool. Because the cocovenenans pathovar is found in such a narrow range of foods, a positive result in this assay using bacteria isolated from a suspect food would be strong evidence for the presence of B. gladioli pv. cocovenenans.

24.2  Methods 24.2.1  Reagents and Equipment Table 24.2 describes the reagents and equipment required for the B. gladioli pv. cocovenenans detection procedure described in Section 24.2.3.

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Table 24.2 List of Reagents and Equipment Required Reagents -Sterile distilled water -10 × PCR buffer -MgCl2 -dNTPs -Primer sets P1–P3 (Section 24.2.3) -Taq polymerase -Agarose -Gel electrophoresis buffer

Equipment -DNA thermal cycler -Gel electrophoresis apparatus -Ultraviolet light box

-Ethidium bromide

24.2.2 Burkholderia gladioli pathovar cocovenenans Sample Collection and Preparation 24.2.2.1  Bacterial Isolation, Propagation, and Storage Because it is the toxins that cause the illness and not the bacteria themselves, B. gladioli pv. cocovenenans isolates are generally collected from the suspect food instead of from the patient (although in some cases they may be collected from vomit).117 These strains can be isolated from food material on a simple nutrient medium such as PDA.56 Other media commonly used for propagation include Luria-Bertani and trypticase soy agar.9 Colonies on PDA appear white or gray and produce a diffusible yellow pigment (toxoflavin).56,64 When the bacteria are grown on CCM, cell numbers reach 1010 within 2 days of incubation at 30–37°C.97 To quantify the bacteria in a sample, 1 g of CCM is blended with 100 ml 0.9% saline solution for 1 min and serial dilutions are made from this mixture.97 Bacteria can be stored at –80°C in 25% (v/v) glycerol.9 24.2.2.2  Toxin Extraction The following procedure for toxoflavin extraction was described by van Damme et al.54 (1) Propagate cells in 2% glycerol, 1% peptone, and 0.5% NaCl or 0.6% KCl 48 h at 28°C. (2) Add a saturating amount of ammonium sulphate at room temperature. (3) Centrifuge 15 min and filter. Extract three times with chloroform (1/2 filtrate volume). (4) Concentrate to 1% of the original culture volume. Centrifuge and filter solution. (5) Add 5/3 filtrate volume of light petroleum. Extract four times with water (1/4 volume). (6) Dilute to 7.5% of the original culture volume. Repeat steps 2 and 3. Evaporate chloroform. (7) Add 1–2% of the original culture volume of n-propanol at 55°C. Filter and incubate at –5°C. (8) The toxoflavin is now crystallized. Wash crystals with propanol and dry ether and dry.

The following procedure for bongkrekic acid extraction was described by Nugteren and Berends.53 (1) Propagate cells in 1 kg modified CCM. (2) Cover with petroleum ether, incubate 1 day at room temperature, and filter. Repeat. (3) Extract with 50 ml aliquots of 2% sodium bicarbonate. (4) Add 2 N H2SO4 until the extract reaches pH 3 and extract with 250 ml peroxide-free ether. Wash 2 × with 100 ml water. Extract with 25 ml aliquots of 2% sodium bicarbonate until no optical activity is measured. This extract (~150 ml volume) contains fatty acids (including capronic and caprylic acids) and approximately 2 g of bongkrekic acid. The bongkrekic acid can then be further purified using a chromatopile apparatus.53

24.2.3 Burkholderia gladioli pathovar cocovenenans Detection Procedure As discussed above, most of the molecular detection procedures developed that recognize B. gladioli pv. cocovenenans are limited by the fact that they simultaneously detect the other pathovars of B. gladioli (and occasionally other species such as B. glumae and B. plantarii) as part of this group. The following procedure, developed by Clode et al., was developed prior to the reclassification of B. cocovenenans as part of B. gladioli.118 In the testing performed by this group, this protocol successfully differentiates the B. gladioli pv. cocovenenans type strain LMG 11626 from the B. gladioli pv. gladioli strains ATCC 25417, ATCC 10247, and type strain ATCC 10248/NCTC 12378, B. glumae strain LMG 1277, and B. plantarii strain LMG 10908. This method uses three sets of primers in a multiplex PCR reaction. These primers are designed to amplify a 323 base pair region of BCC 23S rRNA (primer set P1), a 209 base pair region of BCC 16S rRNA (primer set P2), and a 274 base pair 16S–23S rRNA internal transcribed spacer (ITS) region of B. gladioli (primer set P3). At the time that this paper was written, the BCC was not yet divided into 15 (atleast 17) species, so the primers specific for “B. cepacia” were designed to amplify and tested with the BCC species B. cepacia, B. cenocepacia, B. stabilis, B. vietnamiensis, and B. lata. Primer sequences

Primer set P1 Primer set P2 Primer set P3

PC1 GCTGC GGATG CGTGC TTTGC PC2 GCCTT CTCCA ATGCA GCGAC PSR1 TTTCG AGCAC TCCCG CCTCT CAG PSL1 AACTA GTTGT TGGGG ATTCA TTTC PG1 TTCAA TGACA AACGT TCGGG PG2 GCTTT CGCTT GACAG GCC

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Burkholderia

Multiplex PCR protocol (1) To isolate chromosomal DNA, transfer five colonies (following incubation on solid media for 48 h) to 100 µl sterile distilled water. Vortex and centrifuge 5 min at 13000 × g. Transfer 3 µl of the supernatant to 12 µl sterile distilled water. (2) Set up PCR reactions containing 2.5 µl 10 × PCR buffer, 50 pmol MgCl2, 2.5 mol of each dNTP, 100 pmol of each primer (PC1, PC2, PSR1, PSL1, PG1, and PG2), 1.25 U of Taq polymerase, and the chromosomal DNA prepared in the previous step. (3) Amplify in a DNA thermal cycler under the following conditions: one cycle of 96°C for 5 min; 24 cycles of 96°C for 15 sec, 63°C for 30 sec, 72°C for 90 sec; one cycle of 70°C for 5 min (4) Separate PCR products on a 1.5% (wt/vol) agarose gel at 100 V for 1.5 h. The authors tested this protocol with characterized strains of the BCC (B. cepacia, B. cenocepacia, B. stabilis, B. vietnamiensis, and B. lata), B. gladioli pvs. gladioli and cocovenenans, B. glumae, B. plantarii, Burkholderia andropogonis, Burkholderia caryophylli, Burkholderia vandii, Ralstonia pickettii, Ralstonia solanacearum, and Pseudomonas aeruginosa. Of all of these strains, only the amplification of B. gladioli pv. cocovenenans strain LMG 11626 DNA generated PCR products approximately 274 and 323 base pairs in size (Figure 24.1). Only this DNA was amplified by both the P1 and P3 primer sets, thus differentiating B. gladioli pv. cocovenenans from all other

1.47 Kb 738 bp 369 bp 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21

123 bp

Figure 24.1  Multiplex PCR with three primer sets and reference strains of Burkholderia spp. and other species. Lane 1: B. cepacia NCTC 10661; lane 2: B. cenocepacia NCTC 10744; lane 3: B. cenocepacia ATCC 25610; lane 4: B. gladioli pv. gladioli ATCC 10248/NCTC 12378; lane 5: B. gladioli pv. gladioli ATCC l0248/ NCTC 12378; lane 6: B. gladioli pv. gladioli ATCC 25417; lane 7: R. pickettii NCTC 11149; lane 8: B. gladioli pv. cocovenenans LMG 11626; lane 9: R. solanacearum LMG 2295; lane 10: B. caryophylli LMG 2155; lane 11: B. vandii LMG 16020; lane 12: B. glumae LMG 1277; lane 13: B. vietnamiensis LMG 6998; lane 14: B. plantarii LMG 10908; lane 15: B. andropogonis LMG 2126; lane 16: P. aeruginosa NCTC 10332; lane 17: P. aeruginosa NCTC 10662; lane 18: Acinetobacter baumannii; lane 19: S. maltophilia; lane 20: water blank; lane 21: 123 bp size markers. (From Clode, F.  E. et al., J. Clin. Pathol., 52, 173, 1999. Reproduced/amended with permission from the BMJ Publishing Group.)

Table 24.3 Reactions of Reference Strains of Burkholderia and Ralstonia with Primer Sets Species B. cepacia B. cenocepacia B. vietnamiensis B. cenocepacia B. stabilis B. lata B. cenocepacia B. cenocepacia B. gladioli pv. gladioli B. gladioli pv. gladioli B. gladioli pv. gladioli B. gladioli pv. gladioli B. glumae B. plantarii B. vietnamiensis B. vandii R. solanacearum B. andropogonis B. caryophylli B. gladioli pv. cocovenenans R. pickettii P. aeruginosa

Strain NCTC 10661 NCTC 10744 ATCC 29424 ATCC 25608 ATCC 27515 ATCC 17460 ATCC 25610 ATCC 17765 NCTC 12378 ATCC 25417 ATCC 10247 ATCC 10248 LMG 1277 LMG 10908 LMG 6998 LMG 16020 LMG 2295 LMG 2126 LMG 2155 LMG 11626 NCTC 11149 NCTC 10332

P1*

P2*

P3*

+ + + + + + + + – – – – – – + + + – – + – –

+ + + + + + + + – – – – + – – + – – + – – –

– – – – – – – – + + – + – – – – – – – + – –

*Primers as in Section 24.2.3. Source: Clode, F. E. et al., J. Clin. Pathol., 52, 173, 1999. Reproduced/ amended with permission from the BMJ Publishing Group.

species tested (Table 24.3). When 177 clinical isolates putatively identified as “B. cepacia” were tested with these primer sets, none of them formed products with both P1 and P3, suggesting that reaction with these sets is relatively specific to B. gladioli pv. cocovenenans. Although further testing is required to verify the sensitivity and specificity of the amplification, this protocol appears to effectively identify B. gladioli pv. cocovenenans and differentiate it from B. gladioli pv. gladioli, B. glumae, and B. plantarii.

24.3 Conclusions and Future Perspectives Although B. gladioli pv. cocovenenans is a relatively unknown foodborne pathogen, it causes significant morbidity and mortality in certain parts of the world. There are substantial challenges involved in identifying these bacteria, both with respect to the biology of the organism and the socioeconomic conditions of the affected regions. Appropriate detection methods for B. gladioli pv. cocovenenans should be specific enough to differentiate it from related organisms such as B. glumae, B. plantarii, and the other pathovars of B. gladioli, but should be carried out using relatively inexpensive materials and equipment. Most molecular detection methods developed to date, including CE–SSCP, microarray analysis,

340

and probe-based cell fishing, are deficient in one or both of these areas. Testing performed by Clode et al. suggests that an effective and simple method for the detection of B. gladioli pv. cocovenenans is a multiplex PCR based on amplification of rDNA and internal transcribed spacer sequences.118 However, further studies are required to verify the effectiveness of this protocol. Research into novel diagnostic measures for these organisms, particularly the development of a selective medium, could be instrumental in preventing and responding to future outbreaks and intoxications.

References













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Molecular Detection of Foodborne Pathogens 15. Bezerra Rolim, D. et al. Melioidosis, northeastern Brazil. Emerg. Infect. Dis., 11, 1458, 2005. 16. Whitmore, A. An account of a glanders-like disease occurring in Rangoon. J. Hyg., 13, 1, 1913. 17. Dance, D. A. B. Ecology of Burkholderia pseudomallei and the interactions between environmental Burkholderia spp. and human-animal hosts. Acta Trop., 74, 159, 2000. 18. Suputtamongkol, Y. et al. The epidemiology of melioidosis in urban Ratchatani, northeast Thailand. Int. J. Epidemiol., 23, 1082, 1994. 19. Stanton, A. T. and Fletcher, W. Studies from the Institute for Medical Research, Federated Malay States, No. 21: Melioidosis. John Bale, Sons & Danielsson, Ltd., London, 1932. 20. Inglis, T. J. J. et al. Dry-season outbreak of melioidosis in Western Australia. Lancet, 352, 1600, 1998. 21. Zanetti, F., De Luca, G. and Stampi, S. Recovery of Burkholderia pseudomallei and B. cepacia from drinking water. Int. J. Food. Microbiol., 59, 67, 2000. 22. Rose, L. J. et al. Chlorine inactivation of bacterial bioterrorism agents. Appl. Environ. Microbiol., 71, 566, 2005. 23. Whitlock, G. C., Mark Estes, D. and Torres, A. G. Glanders: off to the races with Burkholderia mallei. FEMS Microbiol. Lett., 277, 115, 2007. 24. Gilad, J. et al. Burkholderia mallei and Burkholderia pseudomallei as bioterrorism agents: national aspects of emergency preparedness. Israel Med. Assoc. J., 9, 499, 2007. 25. Wilson, G. S. and Miles, A. Topley and Wilson’s Principles of Bacteriology, Virology and Immunity, 6th Edition. Edward Arnold, London, 1975. 26. Srinivasan, A. et al. Glanders in a military research microbio­ logist. New Engl. J. Med., 345, 256, 2001. 27. Rotz, L. D. et al. Public health assessment of potential biological terrorism agents. Emerg. Infect. Dis., 8, 225, 2002. 28. Takhistov, P. and Bryant, C. M. Protecting the food supply. Food Technol., 60, 34, 2006. 29. Rosenstein, B. J. and Hall, D. E. Pneumonia and septicemia due to Pseudomonas cepacia in a patient with cystic fibrosis. Johns Hopkins Med. J., 147, 188, 1980. 30. Bottone, E. J. et al. Association of Pseudomonas cepacia with chronic granulomatous disease. J. Clin. Microbiol., 1, 425, 1975. 31. Zhang, L. and Xie, G. Diversity and distribution of Burkholderia cepacia complex in the rhizosphere of rice and maize. FEMS Microbiol. Lett., 266, 231, 2007. 32. Ramette, A. and Tiedje, J. M. Multiscale responses of microbial life to spatial distance and environmental heterogeneity in a patchy ecosystem. Proc. Natl. Acad. Sci. USA, 104, 2761, 2007. 33. Yara, R. et al. A bacterium belonging to the Burkholderia cepacia complex associated with Pleurotus ostreatus. J. Microbiol., 44, 263, 2006. 34. Cohen, R., Persky, L. and Hadar, Y. Biotechnological applications and potential of wood-degrading mushrooms of the genus Pleurotus. Appl. Microbiol. Biot., 58, 582, 2002. 35. Smith, D., Mikolajcik, E. and Lindamood, J. Causative organisms and chemical nature of the Swiss cheese rind rot defect. Cult. Dairy Prod. J., 22, 9, 1987. 36. Blanco, D. et al. Study of “potato defect” of raw dry-cured ham. Ind. Conserv., 69, 230, 1994. 37. Huang, Y. et al. Biocontrol of postharvest orange diseases by a strain of Pseudomonas cepacia under semi-commercial conditions. Postharvest Biol. Tec., 3, 293, 1993.

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341 57. Research Group for Pathogenesis of Fermented Corn Flour Poisoning. A new species of food poisoning bacteria – Flavobacterium farinofermentans sp. nov. Acta Acad. Med. Sincae, 2, 77, 1980. Cited in Zhao, N. X. et al. Comparative description of Pseudomonas cocovenenans (van Damme, Johannes, Cox, and Berends 1960) NCIB 9450T and strains isolated from cases of food poisoning caused by consumption of fermented corn flour in China. Int. J. Syst. Bacteriol., 40, 452, 1990. 58. Zhao, N. X. et al. Comparative study on Pseudomonas cocovenenans and Pseudomonas farinofermentans. Chin. J. Microbiol. Immunol., 8, 151, 1988. 59. Meng, Z. et al. Comparing studies on Flavobacterium farinofermentans sp. nov. (tentative name) and Pseudomonas cocovenenans (NCIB 9450). J. Hyg. Res., 16, 17, 1987. 60. Hu, W. J. et al. Fermented corn flour poisoning in rural areas of China. III. Isolation and identification of main toxin produced by causal microorganisms. Biomed. Environ. Sci., 2, 65, 1989. 61. Zhao, N. X. et al. Comparative description of Pseudomonas cocovenenans (van Damme, Johannes, Cox, and Berends 1960) NCIB 9450T and strains isolated from cases of food poisoning caused by consumption of fermented corn flour in China. Int. J. Syst. Bacteriol., 40, 452, 1990. 62. Yabuuchi, E. et al. Proposal of Burkholderia gen. nov. and transfer of seven species of the genus Pseudomonas homology group II to the new genus, with the type species Burkholderia cepacia (Palleroni and Holmes 1981) comb. nov. Microbiol. Immunol., 36, 1251, 1992. 63. Gillis, M. et al. Polyphasic taxonomy in the genus Burkholderia leading to an emended description of the genus and proposition of Burkholderia vietnamiensis sp. nov. for N2-fixing isolates from rice in Vietnam. Int. J. Syst. Bacteriol., 45, 274, 1995. 64. Zhao, N. et al. Phylogenetic evidence for the transfer of Pseudomonas cocovenenans (van Damme et al. 1960) to the genus Burkholderia as Burkholderia cocovenenans (van Damme et al. 1960) comb. nov. Int. J. Syst. Bacteriol., 45, 600, 1995. 65. Vandamme, P. et al. Occurrence of multiple genomovars of Burkholderia cepacia in cystic fibrosis patients and proposal of Burkholderia multivorans sp. nov. Int. J. Syst. Evol. Microbiol., 47, 1188, 1997. 66. Coenye, T. et al. Burkholderia cocovenenans (van Damme et al. 1960) Gillis et al. 1995 and Burkholderia vandii Urakami et al. 1994 are junior synonyms of Burkholderia gladioli (Severini 1913) Yabuuchi et al. 1993 and Burkholderia plantarii (Azegami et al. 1987) Urakami et al. 1994, respectively. Int. J. Syst. Bacteriol., 49, 37, 1999. 67. Qiu, M., Liu, X. and Yang, R. Study on the molecular epidemiology characteristics of Pseudomonas cocovenenans subsp. farinofermentans isolated from China with rDNA fingerprinting. J. Hyg. Res., 27, 57, 1998. 68. Qiu, M. and Liu, X. Relationship between the toxigenicity and ribotype distribution of Pseudomonas cocovenenans subsp. farinofermentans. J. Hyg. Res., 27, 119, 1998. 69. Jiao, Z. et al. Sequencing and analysis of 16S rDNA sequences for P. cocovenenans subsp farinofermentans. J. Hyg. Res., 28, 232, 1999. 70. Jiao, Z. et al. Determination of DNA-DNA homology among Pseudomonas cocovenenans subsp. farinofermentans with microdilution plate hybridization method. Acta Microbiol. Sinica, 41, 70, 2001. 71. Adams, M. R. and Moss, M. O. Food Microbiology, 2nd Ed. Royal Society of Chemistry, Cambridge, UK, 2000.

342 72. Garcia, R. A., Hotchkiss, J. H. and Steinkraus, K. H. The effect of lipids on bongkrekic (Bongkrek) acid toxin production by Burkholderia cocovenenans in coconut media. Food Addit. Contam., 16, 63, 1999. 73. Jay, J. M., Loessner, M. J. and Golden, D. A. Modern Food Microbiology, 7th Ed., Springer, New York, 2005. 74. van Veen, A. G. The bongkrek toxins, pp. 43–50. In R. I. Mateles and Wogan, G. N. (ed.), Biochemistry of Some Foodborne Microbial Toxins. MIT Press, Cambridge, UK, 1966. 75. Arbianto, P. Bongkrek Food Poisoning in Java. Proc. GIAM-V, 371, 1979. 76. Mujimanto, Memeragi Bongkrek. Topic, 75, 24, 1975. Cited in Arbianto, P. Bongkrek food poisoning in Java. Proc. GIAM-V, 371, 1979. 77. Indonesian Embassy. Tempe, the unique health food of Indonesia [online], http://www.kbri-islamabad.go.id/index. php?option=com_content&task=view&id=30&Itemid=1, 2008. 78. Meng, Z. et al. Studies on fermented corn flour poisoning in rural areas of China. I. Epidemiology, clinical manifestations, and pathology. Biomed. Environ. Sci., 1, 101, 1988. 79. Mahenthiralingam, E., Urban, T A. and Goldberg, J. B. The multifarious, multireplicon Burkholderia cepacia complex. Nat. Rev. Microbiol., 3, 144, 2005. 80. Machlowitz, R. A. et al. Xanthothricin, a new antibiotic. Antibiot. Chemother., 4, 259, 1954. 81. Jeong, Y. et al. Toxoflavin produced by Burkholderia glumae causing rice grain rot is responsible for inducing bacterial wilt in many field crops. Plant Dis., 87, 890, 2003. 82. Sato, Z. et al. Toxins produced by Pseudomonas glumae. Ann. Phytopathol. Soc. Jpn., 55, 353, 1989. Cited in Maeda, Y. et al. Phylogenetic study and multiplex PCR-based detection of Burkholderia plantarii, Burkholderia glumae and Burkholderia gladioli using gyrB and rpoD sequences. Int. J. Syst. Evol. Microbiol., 56, 1031, 2006. 83. Lijmbach, G. W. M., Cox, H. C. and Berends, W. Elucidation of the chemical structure of bongkrekic acid. I. Isolation, purification and properties of bongkrekic acid. Tetrahedron, 26, 5993, 1970. 84. Buckle, K. A. and Kartadarma, E. Inhibition of bongkrek acid and toxoflavin production in tempe bongkrek containing Pseudomonas cocovenenans. J. Appl. Bacteriol., 68, 571, 1990. 85. Daves Jr., G. D., Robins, R. K. and Cheng, C. C. The total synthesis of toxoflavin. J. Am. Chem. Soc., 83, 3904, 1961. 86. Levenberg, B. and Linton, S. N. On the biosynthesis of toxoflavin, an azapteridine antibiotic produced by Pseudomonas cocovenenans. J. Biol. Chem., 241, 846, 1966. 87. Yoneda, F., Shinomura, K. and Nishigaki, S. A convenient synthesis of toxoflavins and toxoflavin-n-oxides. Tetrahedron Lett., 13, 851, 1971. 88. Voet, D., Voet, J. G. and Pratt, C. W. Fundamentals of Biochemistry, Upgrade Edition. John Wiley & Sons, Inc., New York, 2002. 89. Latuasan, H. E. and Berends, W. On the origin of the toxicity of toxoflavin. Biochim. Biophys. Acta, 52, 502, 1961. 90. Lijmbach, G. W. M., Cox, H. C. and Berends, W. Elucidation of the structure of bongkrekic acid. II. Chemical structure of bongkrekic acid and study of the UV, IR, NMR and mass spectra. Tetrahedron, 27, 1839, 1971. 91. de Bruijn, J. et al. The structure of bongkrekic acid. Tetrahedron, 29, 1541, 1973. 92. Hu, W. J., Zhang, G. S. and Chu, F. S. Purification and partial characterization of flavotoxin A. Appl. Environ. Microbiol., 48, 690, 1984.

Molecular Detection of Foodborne Pathogens 93. Welling, W., Cohen, J. A. and Berends, W. Disturbance of oxidative phosphorylation by an antibioticum produced by Pseudomonas cocovenenans. Biochem. Pharmacol., 3, 122, 1960. 94. Henderson, P. J. and Lardy, H. A. Bongkrekic acid: an inhibitor of the adenine nucleotide translocase of mitochondria. J. Biol. Chem., 245, 1319, 1970. 95. Buchanan, B. B., Eiermann, W. and Riccio, P. Antibody evidence for different conformational states of ADP,ATP translocator protein isolated from mitochondria. Proc. Natl. Acad. Sci. USA, 73, 2280, 1976. 96. Schwerdt, G. et al. Inhibition of mitochondria prevents cell death in kidney epithelial cells by intra- and extracellular acidification. Kidney Int., 63, 1725, 2003. 97. Ko, S. D. Growth and toxin production of Pseudomonas cocovenenans, the so-called ‘bongkrek bacteria’. ASEAN Food J., 1, 78, 1985. 98. Ko, S. D., Kelholt, A. J. and Kampelmacher, E. H. Inhibition of toxin production in tempe bongkrek. Proc. GIAM-V, 375, 1979. 99. Segonds, C. et al. Differentiation of Burkholderia species by PCR-restriction fragment length polymorphism analysis of the 16S rRNA gene and application to cystic fibrosis isolates. J. Clin. Microbiol., 37, 2201, 1999. 100. Voragen, A. G. J., De Kok, H. A. M. and Kelholt, A. J. Determination of bongkrek acid and toxoflavin by high pressure liquid chromatography. Food Chem., 9, 167, 1982. 101. Coenye, T. et al. Taxonomy and identification of the Burkholderia cepacia complex. J. Clin. Microbiol., 39, 3427, 2001. 102. Bauernfeind, A. et al. Molecular procedure for rapid detection of Burkholderia mallei and Burkholderia pseudomallei. J. Clin. Microbiol., 36, 2737, 1998. 103. Viallard, V. et al. Burkholderia graminis sp. nov., a rhizospheric Burkholderia species, and reassessment of [Pseudomonas] phenazinium, [Pseudomonas] pyrrocinia and [Pseudomonas] glathei as Burkholderia. Int. J. Syst. Bacteriol., 48, 549, 1998. 104. Ferroni, A. et al. Use of 16S rRNA gene sequencing for identification of nonfermenting gram-negative bacilli recovered from patients attending a single cystic fibrosis center. J. Clin. Microbiol., 40, 3793, 2002. 105. Boyanton Jr., B. L. et al. Burkholderia gladioli osteomyelitis in association with chronic granulomatous disease: Case report and review. Pediatr. Infect. Dis. J., 24, 837, 2005. 106. Lynch, K. H. and Dennis, J. J. Development of a species-specific fur gene-based method for identification of the Burkholderia cepacia complex. J. Clin. Microbiol., 46, 447, 2008. 107. Höflich, C. et al. Naturally occurring anti-IFN-γ autoantibody and severe infections with Mycobacterium cheloneae and Burkholderia cocovenenans. Blood, 103, 673, 2004. 108. Halle, E. et al. Melioidosis-like disease: infection with Burkholderia cocovenenans as rare differential diagnosis for lymphadenitis colli. Laryngo-Rhino-Otol., 86, 287, 2007. 109. Ghozzi, R. et al. Capillary electrophoresis-single-strand conformation polymorphism analysis for rapid identification of Pseudomonas aeruginosa and other Gram-negative nonfermenting bacilli recovered from patients with cystic fibrosis. J. Clin. Microbiol., 37, 3374, 1999. 110. Jin, L. Q. et al. Detection and identification of intestinal pathogenic bacteria by hybridization to oligonucleotide microarrays. World J. Gastroentero., 11, 7615, 2005. 111. Wang, X. W. et al. Development and application of an oligonucleotide microarray for the detection of food-borne bacterial pathogens. Appl. Microbiol. Biot., 76, 225, 2007.

Burkholderia 112. Stoffels, M., Ludwig, W. and Schleifer, K. H. rRNA ­probe-based cell fishing of bacteria. Environ. Microbiol., 1, 259, 1999. 113. Brisse, S. et al. Distinguishing species of the Burkholderia cepacia complex and Burkholderia gladioli by automated ribotyping. J. Clin. Microbiol., 38, 1876, 2000. 114. Whitby, P. W. et al. Species-specific PCR as a tool for the identification of Burkholderia gladioli. J. Clin. Microbiol., 38, 282, 2000. 115. Furuya, N. et al. Specific oligonucleotide primers based on sequences of the 16S-23S rDNA spacer region for the detection of Burkholderia gladioli by PCR. J. Gen. Plant Pathol., 68, 220, 2002.

343 116. Maeda, Y. et al. Phylogenetic study and multiplex PCR-based detection of Burkholderia plantarii, Burkholderia glumae and Burkholderia gladioli using gyrB and rpoD sequences. Int. J. Syst. Evol. Microbiol., 56, 1031, 2006. 117. Jakarta Post. Burkholderia cocovenenans foodborne illness – Indonesia (Central Java) [online], http://www.promedmail. org/pls/otn/f?p=2400:1202:3714406431165003::NO::F240 0P1202_CHECK_DISPLAY,F2400_P1202_PUB_MAIL_ ID:X,38630, 2007. 118. Clode, F. E. et al. Evaluation of three oligonucleotide primer sets in PCR for the identification of Burkholderia cepacia and their differentiation from Burkholderia gladioli. J. Clin. Pathol., 52, 173, 1999.

25 Campylobacter

Aurora Fernández Astorga and Rodrigo Alonso Basque Country University

Contents 25.1 Introduction.................................................................................................................................................................... 345 25.1.1 Classification..................................................................................................................................................... 345 25.1.2 Biology.............................................................................................................................................................. 346 25.1.3 Pathogenesis...................................................................................................................................................... 346 25.1.3.1 Virulence Factors............................................................................................................................. 346 25.1.3.2 Animal Models................................................................................................................................ 348 25.1.4 Isolation and Detection..................................................................................................................................... 348 25.1.4.1 Traditional Methods......................................................................................................................... 348 25.1.4.2 Rapid Methods................................................................................................................................. 349 25.1.4.3 Molecular Methods.......................................................................................................................... 349 25.2 Methods.......................................................................................................................................................................... 353 25.2.1 Reagents and Equipment................................................................................................................................... 353 25.2.2 Sample Preparation and DNA Extraction......................................................................................................... 353 25.2.3 Detection Procedures........................................................................................................................................ 354 25.2.3.1 Multiplex PCR................................................................................................................................. 354 25.2.3.2 Duplex Real-Time PCR................................................................................................................... 355 25.3 Conclusion and Future Perspectives............................................................................................................................... 355 Acknowledgments...................................................................................................................................................................... 356 References.................................................................................................................................................................................. 356

25.1  Introduction 25.1.1  Classification Members of the genus Campylobacter are classified into the epsilon class of proteobacteria, in the order Campylobacteriales. They share a close taxonomic framework and biological and clinical characteristics which complicates the species-specific identification in some cases. The genus Campylobacter, first described by Véron and Chatelain1 in 1973, comprises Gram-negative, nonspore-forming bacilli that have a curved or spiral shape with tapering ends. Cells possess a polar flagellum at one or both ends of the cells which is responsible of the characteristic corkscrew-like motility. They are generally microaerophilic requiring modified atmospheres with 3–15% oxygen and 2–10% carbon dioxide for optimal growth conditions. Further, they are unable to ferment nor to oxidize carbohydrates and to degrade complex substances instead they obtain energy from amino acids, or tricarboxylic acid cycle intermediates.2,3 At present, the genus includes at least 16 species, and six subspecies, of which Campylobacter jejuni subsp. jejuni, C. jejuni subsp. doylei, Campylobacter coli, Campylobacter lari, Campylobacter upsaliensis, and Campylobacter helveticus form a genetically closed group of species4 classed as thermophilic owing its optimal growth at 42°C. Thermophilic

campylobacters infect both human and warm-blooded animals. Although primarily occurring as commensals in a wide range of domestic and wild birds and animals, thermophilic campylobacters (except C. jejuni subsp. doylei) can also cause diseases being the most commonly isolated from human and animal diarrhea.4–6 As animal pathogens they are long related to diarrhea in cattle and septic abortion in both cattle and sheep,7 however, it was not until the 1970s that they were successfully associated to human enteritis (for an historical account, see Skirrow8). Human infection is termed campylobacteriosis, and ranges from self-limiting gastroenteritis to more serious systemic infections, i.e., bacteriaemia or hemolytic uraemic syndrome.6,9 Post-infectious complications are rare but demyelinating disorders as Guillian–Barré syndrome (GBS) and Miller Fisher syndrome (MFS) are now recognized as sequelae of Campylobacter infection.10 The epidemiology of Campylobacter infection is complex since the organism is widely distributed in the environment and throughout the food chain. These human pathogens are commonly found in the environment and on many raw foods, of both plant and animal origin. Campylobacters are considered to be part of the normal intestinal biota of a wide range of domestic and wild animals. Fecal contamination of meat often occurs during slaughtering, and human infection 345

346

is usually acquired through the consumption of undercooked contaminated meat or other cross-contaminated food products. Bacterial numbers can be very high on certain key foods like raw poultry meat. The risks to human health vary between the different animal species and will also be different between countries often due to variations in food preparation and consumption patterns.6 With incidences in developed countries two to seven times more frequent than infections with Salmonella species, Shigella species, or Escherichia coli O157:H79 campylobacteriosis is now considered to be one of the most important foodborne diseases worldwide.6 A range of food of mainly animal origin is involved in human transmission but it is of general acceptance that consumption and handling of poultry11 are the major sources of Campylobacter infection even though other sources like unpasteurized milk,12,13 contaminated water,14,15 and contact with pets especially birds and cats16 are also common. Although some outbreaks have been reported17–19 most campylobacteriosis cases are normally sporadic, affecting children and young adults in developed countries and mainly infants and young children in developing countries.9 Owing to the incidence, thought to be an estimated 1% of the population per year in developed countries and even higher in developing countries, as well as the spectrum of disease with chronic sequelae and the food risks, it is not surprising that the health and social burden of campylobacteriosis could be larger than estimated.20 Among the etiological agents of human campylobacteriosis it is worth highlighting C. jejuni which accounts for more than 95% of bacteria isolated from diarrheal stool samples.21–23 Therefore C. jejuni is a pathogen of considerable clinical and economic importance and requires easy and rapid detection to facilitate precautionary food safety measures. The conventional microbiological methods, however, usually include multiple subcultures and biotype or serotype-identification steps and, thus are laborious and timeconsuming. As a consequence, this chapter concentrates on both well-established aspects and recent advances in the characterization of C. jejuni and the alternative methods for its identification in food.

25.1.2  Biology C. jejuni was first described by Jones et al.24 in 1931, who named it Vibrio jejuni because of its similarity to so-called V. fetus (current Campylobacter fetus) found in aborting cattle.7 Within the species C. jejuni two subspecies, ssp. doylei and ssp. jejuni, can be distinguished on the basis of nitrate reduction, cephalothin susceptibility and catalase activity. With the pathogenic role of the ssp. doyley yet not defined,3 hereafter in this chapter C. jejuni will refer to C. jejuni ssp. jejuni. C. jejuni cells have a classic spiral morphology, 0.2–0.8 µm wide and 0.5–5.0 µm long. In old cultures or under stressed conditions they tend to spherical morphology (coccoid forms) with a diameter of approximately 1 µm. Cells are motile by means of unipolar or bipolar unsheathed flagella. The insertion of the flagellum in a cone-like depression allows its

Molecular Detection of Foodborne Pathogens

rotation in a characteristic corkscrew-like manner providing an extremely rapid, spinning motion to the cell.25 Due to this ability C. jejuni can move in very viscous media and can easily colonize and pass through the mucous layer in the intestinal tract of humans and animals.26 The genome of C. jejuni is a 1.6–1.7-Mbp adenine and thymine AT-rich DNA and contains a low G + C ratio, 30 mol% on average.27,28 This is a relatively small genome compared with other enteropathogens such as Escherichia coli (4.5 Mbp), and Salmonella enterica (4.9 Mbp). Some of the remarkable biochemical characteristics, i.e., their inability to ferment nor to oxidize carbohydrates and to degrade complex substances as well as their requirement for complex media for growth, stem from the small size of the genome.28 The small size of the genome has facilitated additional genome sequencing since the first NCTC 11168 C. jejuni genome sequence data were available in 2000.29 At present, a total of eight C. jejuni whole-genomes are available on the website of the National Center for Biotechnology Information (NCBI), http://www.ncbi.nlm.nih.gov. Of note are the hypervariable regions found in the C. jejuni genome which might be important both in survival30 and pathogenesis of the organisms.31,32 The genome sequence, however, contains no transposons, phage remnants, or insertion sequence elements and very few repeat sequences,29 making C. jejuni almost unique among sequenced bacterial pathogens.33 Typically microaerophilic and moderately capnophilic, the organisms require lower O2 and higher CO2 concentrations than the ambient atmosphere and may be cultured in atmospheres with 5–10% O2 supplemented with 2–10% CO2. However, C. jejuni also grows fairly well in humidified CO2 incubators ranging from 5 to 14% CO2 as claimed by Gaynor et al.34 The range of growth temperatures is narrow from 30 to 44ºC, with an optimal temperature of 37 and 42ºC. Even at optimal conditions the growth is quite slow and subsequently the times for primary isolation are long, 2–5 days.

25.1.3  Pathogenesis A reasonable understanding of the general clinical, microbiological, and epidemiological aspects of Campylobacter infection has been achieved. However, the molecular mechanisms involved in pathogenesis are still poorly understood. Few of the virulence determinants involved in Campylobacter pathogenesis are known or have a proven role. The virulence factors are generally not well characterized and some are rather controversial. In association with food or water, campylobacters enter the host intestine via the stomach acid barrier and colonize the distal ileum and colon. After colonization of the mucus and adhesion to intestinal cell surfaces, campylobacters perturb the absorptive capacity of the intestine by damaging epithelial cell function. 25.1.3.1  Virulence Factors Chemotaxis and motility. Effective colonization requires chemotaxis. Genome sequence analysis has shown that the C. jejuni genome encodes most features of the E. coli chemotaxis

Campylobacter

system.29,35 C. jejuni displays chemotactic motility towards mucin, L-serin, and L-fucose, amino acids that are found in the chicken gastrointestinal tract and mucus components.36 Mutants which lack either of the chemotaxis receptors DocB or Cj0262c show decreased chick colonization.37 Several other components of the chemotaxis system of C. jejuni have been identified, including CheY, CheV, CetA, and CetB.38 Motility of Campylobacter spp. necessitates the production of flagellum, the best characterized virulence determinant of campylobacters. Flagella and flagellar motility are vital to host colonization, virulence in ferret models, secretion, and host-cell invasion.38 The flagella of C. jejuni consists of an unsheathed polymer of flagellin subunits, which are encoded by the adjacent flaA and flaB genes. Both genes are subjected to antigenic and phase variation. Mutants of flaA, the primary structural gene for flagella, are unable to colonize chicks and cannot invade human intestinal epithelial cells in vitro.39 Adhesion and invasion are dependent on both motility and flagella expression, as C. jejuni mutants with reduced motility show reduced adherence and no invasion. This indicates that, while flagella are involved in adherence, other adhesins are involved in subsequent internalization.40 Adhesion and invasion. Adhesion by bacterial pathogens is often mediated by fimbrial structures. Genome annotations of several C. jejuni strains do not include obvious pilus or pilus-like ORFs. However, several proteins contribute to C. jejuni adherence to eukaryotic cells. CadF mediates adhesion by binding to the cell matrix protein fibronectin.41 CadF is required for maximal binding and invasion by C. jejuni in vitro, and cadF mutants are unable to bind fibronectin and did not colonize chicks.42 Another characterized adhesin, JlpA, is a surface exposed lipoprotein that has been shown to bind to Hsp90α on Hep-2 cells. This binding resulted in activation of NF-κB and p38 mitogen-activated protein kinase, both of which contribute to the inflammatory response.43,44 The lipoprotein CapA, a putative autotransporter, has been described as a possible adhesin that plays a role in adhesion and invasion of Caco-2 cells. Additionally, CapA-deficient mutants have decreased colonization and persistence in a chick model.45 Another reported adhesin is Peb1, a periplasmic ABC binding protein that is required for adherence to HeLa cells and for intestinal colonization of mice.46 C. jejuni is generally considered to be invasive,47 although the mechanism of uptake into epithelial cells remains vague. There are no specialized type III secretion systems similar to those of other enteric pathogens that mediate invasion into epithelial cells. Instead, the flagella appears to function as a type III secretion organelle that secretes a number of proteins, some of which are reported to affect invasion. The Cia proteins (for Campylobacter invasion antigens) are secreted through the flagella filament and affect invasion of some strains of C. jejuni.48,49 Although CiaB is internalized into epithelial cells, the mechanism by which CiaB mediates invasion has not been detailed.32 CiaB and other secreted Cia proteins (Cia A–H) require a functional flagellar export apparatus for their secretion.49 As well as the Cia proteins, this flagellar export apparatus secretes FlaC, which is also required for

347

invasion. Thus, the flagellar export apparatus is an ­important secretion mechanism in C. jejuni and is required for hostcell invasion.38 The ability to cross the epithelial cell barrier either through epithelial cell invasion or via tight junctions, allows the bacterium to move to the basolateral surface. At this point, it can reinvade the epithelial cell or be taken up into macrophages.32 It has been shown that C. jejuni can replicate intracellularly in macrophages and induce apoptosis.50 Interactions of C. jejuni with epithelial cells, dendritic cells and macrophages can release chemokines and cytokines that contribute to both inflammatory diarrhea and clearance of infection.32 The exact mechanisms by which C. jejuni induces disease remains unknown; symptoms could be a result of cytolethal distending toxin (CDT)-induced host cell death and subsequent inflammatory responses.51 CDT. The best characterized virulence factor present in the genome of C. jejuni is CDT. CDT consists of three subunits (CdtA, -B, and -C) encoded by a three-gene operon cdtABC,52 and isogenic C. jejuni cdt mutants lost all CDT activity.53 The role of CDT in C. jejuni pathogenesis remains unclear, but its mechanism of action is becoming understood.38 CDT is a tripartite toxin in which CdtB is the active subunit and CdtA and CdtC comprise the binding components. Once the toxin is bound to the cell, CdtB is transported to the nucleus where it acts as DNase and arrests the cell in the G2 phase of the cell cycle.54 CDT activity causes certain cell types (such as HeLa cells and Caco-2 cells) to become slowly distended, which progresses into cell death.40 Also, CDT induces IL-8 secretion from epithelial cells, which would contribute to inflammatory diarrhea.55 Although all C. jejuni strains tested contain cdt genes, there is a profound variation in CDT titers.56,57 Glycosylation system. Recently, the long-accepted dogma that bacteria only express nonglycosylated protein has been disproved.52,58 In 1989, Logan et al. provided clear evidence that Campylobacter flagellin was post-translationally modified.59 Today it is known that Campylobacter contains both a general N-linked protein glycosylation pathway (responsible for post-translational modification on at least 30 proteins) and an O-linked system (responsible for flagellar glycosylation).60 Both systems have been involved in C. jejuni virulence. Studies about the biological role for N-linked glycosylation clearly links it to bacterial virulence; a pglH mutant showed reduced ability to adhere/invade human epithelia and colonize chicks.61 It has been shown that O-linked glycosylation is essential for successful flagellin assembly and motility hence influencing adhesion, invasion, and virulence in vivo.62 Capsule and lipooligosaccharide (LOS). Genome sequences and DNA microarray data have demonstrated that C. jejuni genome contains about 22 variable regions on the chromosome. The most variable chromosomal regions are those involved in biosynthesis of surface carbohydrate structures, all of which play a role in virulence: the LOS core, a polysaccharide capsule (CPS), and the locus for O-linked glycosilation.32 The C. jejuni polysaccharide capsule is important for serum resistance, the adherence and invasion of

348

epithelial cells, chick colonization and virulence.63 Another study utilizing a capsule-deficient mutant suggested that contribution of bacterial CPS to host-pathogen interactions may be species-dependent.64 The C. jejuni LOS are highly variable due to differences in monosaccharide linkage and composition. Various LOS structures resemble human neuronal gangliosides. This molecular mimicry is thought to lead to autoimmune disorders, including GBS and MFS syndromes.38 The C. jejuni LOS and flagella have been shown to be sialylated, which is thought to be responsible for the ganglioside mimicry leading to GBS.40,65 Mutations in the various genes that are involved in LOS biosynthesis affect serum resistance, and adherence/ invasion of INT 407 cells.66 25.1.3.2  Animal Models Despite its global importance and the progress that has been made in recent years, there are still large gaps in our knowledge on basic aspects of the mechanisms of C. jejuni pathogenicity and host responses to infection.38 Many questions regarding its biology and pathogenic properties remain unanswered, in particular it is still unclear how C. jejuni causes diarrhea.32,38,67 One of the major obstacles to solve these problems is the lack of a suitable small animal model of infection that closely mimics human disease.68,69 We can find many references in the literature about the use of animal models for C. jejuni infection, but all of them have major disadvantages in terms of disease reproducibility or inadequate biological characterization. Nonhuman primates,70 mice,71,72 ferrets,73 piglets,74 hamsters,75 rabbits76, and chickens77 have been used as models for C. jejuni pathogenesis. Chicken models of commensal infection, mainly chick models, are of interest for the extensive and asymptomatic colonization of the gut. However, these models are limited by the poorly defined immune system of the chick and its unsuitability for genetic manipulation, thereby limiting the understanding of the enteric infection mechanism.67,69 They are used for investigations of the colonization factors with wild and mutant strains66 as well as for testing vaccine efficacy.78–80 Recently new and promising murine models have been developed. Using mice with limited enteric biota81 or immunodeficient mice82–84 some of these limitations can be overcome. Mice, in these models, develop colonization, severe inflammation in the lower intestine and pathological lesions similar to those reported in humans. In spite of many attempts, there is as yet no particular model as suitable for the study of Campylobacter infection in mammals. Other than animal models, polarized intestinal epithelial lines85 as well as human intestinal tissue86 are also used to investigate the pathogenic properties of C. jejuni and the immune response of host. They include INT 407,55 Caco-2,87 and T84 cells,88 and ex vivo organ culture.89

25.1.4  Isolation and Detection 25.1.4.1  Traditional Methods The microbiological analyses of a food are critical to determine the presence of any pathogen organism in order to

Molecular Detection of Foodborne Pathogens

ensure the safety of consumers. Food samples must be ­carefully examined in terms of the isolation of foodborne pathogens. Both false positive (isolated organisms which are not responsible of the specific illness) and false negative (unfounded organisms which are responsible for the specific illness) are important problems in microbiological food analysis. The complex matrix together with the high levels of commensal microbiota in the food may represent larger challenges than just their isolation in clinical samples. Pathogens in clinical samples are normally present as major or unusual microbiota. In contrast, the target microorganism in foods may be present in very low numbers or may be seriously injured by processing and conservation procedures such as freezing, cooling, heating, and salting.90 Even with improved culture methods it could be very difficult to detect these small numbers of foodborne pathogens amid large numbers of indigenous microbiota in a complex sample matrix like foods. The conventional methods based on culture techniques, although very valuable and constantly improving, are both time and material consuming as well as laborious and may be unsuitable in outbreak investigations and for positive release. Positive release involves testing the products to show that they are pathogen-free before they are put in the food chain.6 With some exceptions, the conventional microbiological methods for detection of bacteria in food samples usually include multiple subcultures and biotype or serotypeidentification steps. The presumptive detection requires an overnight pre-enrichment before the enrichment period of 24–48 h, followed by an additional 24–48 h incubation in selective agar plates. Afterwards, the colonies showing an expected morphology must be confirmed, species differentiated and sometimes subtyped by phenotypic characterization based mainly on biochemical and immunological tests. With species identification relying on just a few biochemical tests, in many cases inconclusive, it is not surprising that these conventional methods may sometimes lead to equivocal results and errors in taxonomy. All of that is applicable for the practical totality of the foodborne bacterial pathogens but it accounts especially for those considered fastidious owing the slow growth, susceptibility to environmental factors and restrictive culture conditions such as C. jejuni. The media that are used for isolating C. jejuni from foods are derived from those originally designed for isolating the pathogen from human stool samples.91–94 These media allow the recovery of C. jejuni from fecal samples by direct plating onto selective media. However, due to the predicted low numbers present in foods, selective enrichment broths are also required for the isolation of C. jejuni in foods. Thus, the time to confirm the presence of C. jejuni in food samples can exceed five working days. A standard procedure for the isolation of the organisms includes a pre-enrichment period in a selective broth for 4–6 h at 37ºC to promote the recuperation of the sub-lethally damaged cells; and then an enrichment period in the same medium for 42–44 h at 42ºC to promote specific growth. Afterwards, adequate dilutions of the culture must be spread onto selective agar plates and incubated for an additional 48-h period at 42ºC. Confirmation of the suspected

Campylobacter

colonies can be achieved by minimal standards which are colony morphology, Gram stain, motility, and oxidase test.95 Isolation of the colonies in separate plates requires additional incubations in selective agar plates for species differentiation and typing. All incubations must be under microaerophilic conditions using microaerobic-atmosphere-generating systems. Both solid and liquid selective media are normally achieved by using antibiotics. Typically, these are cefoperazone, cycloheximide, trimethoprim, rifampicin, vancomycin, polymyxin B, and amphotericin or nystatin, combined in different manners.96 Most current media use a combination of cefoperazone and amphotericin B, active against Grampositive bacteria and fungi respectively, together with vancomycin or trimethoprim, but depending on the food examined this cocktail can vary. In addition, selective media usually contain sheep or horse sterile lysed blood to neutralize the toxic effects of compounds formed in the presence of light and oxygen. The need to use lysed blood on media is another critical issue of the traditional methods to be applied in food analyses. It is expensive, easily contaminated and cannot be stored for long periods at the laboratory. Instead of, or in addition to, the lysed blood, charcoal, hematin, and a combination of ferrous sulphate, sodium metabisulfite and sodium pyruvate (FBP supplement) can be used as oxygenquenching ingredients. Attempts to improve oxygen protection include lysed blood and FBP supplement in the same selective media.96 There is little consensus on methods for isolating Campylobacter from foods and water and a number of different isolation methods are used worldwide. However, as with other foodborne bacterial pathogens, it is likely that no single method is ideal for the entire range of foods requiring testing. The 8th edition of the FDA Bacteriological Analytical Manual97 and the online revisions at http://www.foodinfonet. com/publication/fdaBAM.htm give a detailed description of the isolation of Campylobacter from food and water. Data suggest that Bolton broth as enrichment and the modified blood-free mCCDA medium as selective agar give the highest isolation rates for C. jejuni.98 Identification of Campylobacter species is based on biochemical tests such as nitrate/nitrite reduction, hippurate hydrolysis, catalase, and oxidase tests. However, the genus include closely related bacteria that share many biochemical characteristics which makes species differentiation difficult. By way of an example, the hippurate hydrolysis is thought only positive in C. jejuni.99 However, some C. jejuni isolates are hippuricase-negative, making it impossible to differentiate C. coli from hippuricase-negative C. jejuni using purely biochemical tests.39,100 In addition to the inherent difficulties in the detection of C. jejuni such as they may be lost among a background of indigenous microbiota, there is the fact that it can enter a viable but nonculturable state (VBNC) under adverse conditions. The former can be minimized by using pre-enrichment periods but VBNC cells are living cells unable to grow or divide in these media.101 Since ability to enter the VBNC

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state in C. jejuni cells was first described by Rollins and Colwell,102 a number of studies have been developed in order to explain the nature of such cells and the potential public health hazards they present. It has been found that, as culturable cells, they retain spiral morphology103 and metabolic activity.104,105 It has been also suggested that this dormant-like state is an adaptation to survival in adverse environments not supporting growth of Campylobacter, as during transmission or storage.102,106 However, the significance in transmission of disease is today uncertain, as the resuscitation back to the actively growing state of VBNC-forms of Campylobacter cells is still controversial. While several authors have reported C. jejuni resuscitation from the VBNC state after passage in embryonated eggs107 or experimental animals,108–110 there are others who referred their inability to resuscitate.111–113 The resuscitation of the VBNC cells is a key issue for considering the presence of these forms as true health hazards when found in food or water. Owing to the difficulties in finding an adequate animal model for studying the C. jejuni pathogenesis, the differences on resuscitation issues can be explained on the basis of this factor as well as derived for the different strains used in the studies. Although not conclusive, these results show that the presence of C. jejuni VBNC cells in food is of special concern, at least for certain strains, because traditional culture methods cannot elucidate. 25.1.4.2  Rapid Methods From a public health perspective, faster detection times are essential to prevent the spread of infectious diseases or the identification of a continuing source of infection. With the implementation of the HACCP system for control of the process line at CCPs (Critical Control Points), the demand for rapid, sensitive, and accurate methods to detect biological contaminants has also increased. In the past two decades, therefore, large efforts have been made to develop methods for ‘rapid diagnoses’ of foodborne pathogens. Rapid diagnoses include a wide range of novel testing procedures which can significantly reduce the reporting time compared with conventional bacterial culture.114 Following advances in molecular biology techniques and technologies, rapid methods such as antibody-based tests,115,116 simple miniaturized biochemical assays, physicochemical tests (e.g., Fourier transform infrared spectroscopy),117,118 and highly specific nucleic acid-based methods,119–124 have been developed for the specific detection, identification, and typing of microorganisms. The advent of whole-genome based methods, such as DNA microarrays, has generated new opportunities to detect pathogens in foods.125 In particular, methods targeting nucleic acids, both DNA and RNA, are intrinsically more precise and less affected by natural variation than the phenotypic methods. An extensive overview of genetic methods affecting detection and identification of C. jejuni in foods is presented below; the assay principles and some of the detailed procedures are also discussed. 25.1.4.3  Molecular Methods (i) DNA-based methods: DNA-based methods use PCR as the most versatile and widely used amplification technique.

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PCR is an in vitro enzymatic amplification of DNA or RNA, that requires a DNA template, short DNA fragments or primers (usually 20–30 nucleotides long), a thermo stable DNA polymerase like Taq polymerase and a thermocycler. The amplification is directed at short regions of the genome where the primers bind to their complementary bases. It is a cyclical process temporary divided in three thermal steps: (i) denaturization of the DNA template, accomplished by heating to 90–96ºC to open the double-strand of the template into single-strand; (ii) annealing of the primers to their complementary bases on the single-strand template; (iii) and DNA synthesis by the thermostable DNA polymerase to extend the DNA product. The whole process is repeated 25–30 times (cycles) so that within a short period (usually 1 h) there is enough DNA target to detect. A single copy of DNA template can turn into billions of copies in less than 2 h. The products of PCR are normally separated by electrophoresis in agarose gel and detected by a DNA stain. Alternatively, it can be detected by hybridization with labeled probes126 or nucleotide sequencing analysis.127 The hazards of cross-contamination with the PCR products have promoted the development and wide use of the realtime PCR.128 This technique combines PCR methodology with hybridization allowing the amplification and the detection of the products in a single reaction vessel. Detection is based either on fluorescent labeled probes or fluorescent dyes as SYBR green both binding to the PCR products as fast as they are synthesized by the DNA polymerase. The detection is achieved by a fluorimether combined with the thermocycler and recorded during the exponential phase of the amplification without post-PCR handling. This allows for direct detection and quantification as the number of cycles required to reach the threshold value can be directly correlated with the initial numbers of cells in the sample. In comparison, real-time PCR is faster than conventional PCR and may be carried out in automated devices that require less expertise.114 Both techniques can be used as multiplex PCR by including various primer sets within a single reaction. Multiplex PCR may be useful for identification of the pathogens by combining genus- and species-specific genes as targets129–131 as well as in epidemiological studies where a wide range of items may be analyzed.132 However, the selection of the primers should be done carefully. All of them must have similar annealing temperatures, absence of complementarity and enough difference in size for ensuring electrophoresis separation and accurate detection. Regarding C. jejuni, there is a large variety of PCR assays proposed to detect and to identify this species. Most of them are assayed with pure cultures or are developed for human stool samples. In contrast, the range of the applications of these methods for direct detection of C. jejuni in foods is narrow. A summarized description of some of those applied to C. jejuni detection in foods is given in Table 25.1. The consensus major advantages of the PCR-based detection is the speed, together with high specificity and sensibility. One of the main disadvantages, is the presence of false negative assay due to the presence of inhibitory substances

Molecular Detection of Foodborne Pathogens

in the food products which can affect primer binding or Taq ­activity. Part of this undesirable effect reflects the need for a short enrichment period, as much of the methods include, but also reducing the extreme sensitivity of the PCR reaction determined by pure culture when testing foods. Isolation of the DNA to be used in the amplification by in-house methods or commercial kits also minimizes this limitation. The selection of the genetic target to be amplified is one of the more critical aspects when designing a PCR method. As PCR selectively amplifies targets, depending on the specificity, different regions of the genome may be amplified. This is shown in the variety of gene targets used in the PCR-based methods for the detection of C. jejuni. The rRNA presents highly conserved regions in the species of the genus, which is used for Campylobacter spp. detection. In addition, the high numbers of copies per cell provide a naturally amplified target thus increasing the analytic accuracy. The 16S rRNA genes are the most suitable for this purpose and so they are widely used for Campylobacter spp. detection. However, not all the PCRs targeting the 16S rRNA are directed to the same position into the gene sequence. Other genetic targets are selected amid genes exclusively found in a particular species. Such genes are used for differentiating C. jejuni from the other species in the genus. An example of this is the hippurate gene, hipO, coding for the hippurate hydrolase which is found exclusively in C. jejuni species. Since hippurate negative strains were detected, this gene is no longer used in the identification of C. jejuni. Instead, interest has turned to the species-specific fragments of common genes suitable for discriminating between very closed related C. jejuni and C. coli. As an example, the ceuE gene coding a virulence determinant is present in both species but by choosing adequate primers, a specific fragment for each one may be obtained.163 The primers described by Gonzalez et al., for C. coli identification, are used later by a number of authors within different PCR methods.131,156,164 As a result, the test methods and primers used are heterogeneous, and validation and optimization of in-house assays requires further study.114 A major disadvantage concerning the DNA-based methods is the question whether or not they are detecting DNA from dead cells. Due to the long persistence of DNA in cells post-death, the correlation between presence of DNA and viability is not cleared.165 However, this is irrelavant for those methods using a pre-enrichment period. (ii) RNA-based methods: The most commonly used techniques for RNA amplification are reverse PCR (RT-PCR) and nucleic acid sequence-based amplification (NASBA). Both of these have been used to determinate the viability of the molecular targets165 but only NASBA has been also used for Campylobacter detection in food (see Table 25.1). The NASBA technique is a cycling transcriptional process in which single-stranded RNA sequences are targeted and amplified.166 The amplification involves the simultaneous use of three enzymes which mimic the retroviral replication, in a one-step process. Enzymes are, an avian myeloblastosis virus reverse transcriptase (AMV-RT), RNase H, and

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Campylobacter

Table 25.1 Identification of Campylobacter Species by Molecular Methods Target

Matrix

C. jejuni (cadF)

Chicken rinses

C. jejuni (ORF-C sequence)

Raw chicken, offal, shellfish, raw meat, milk Chicken rinses

Thermotolerant campylobacters (16S rRNA) C. jejuni (cadF)

Isolation Procedure Real-Time PCR SE (Mueller-Hinton Broth containing cefoperazone and growth supplements) SE (Bolton broth)

PCR Product Detection

Reference

Melting peak análisis

133

Taqman probe

134

SE (Bolton broth)

Taqman probe

135

Chicken skin

SE (Bolton broth)

Melting peak análisis

136

C. jejuni, C. coli, C. lari (16Sr RNA); C. jejuni (hipO)

Chicken samples

SE (Preston broth)

137

C. jejuni (cadF)

Chicken skin, pork, milk

SE (Bolton broth)

Lightcycler probe (16S rRNA) and melting peak analysis (hipO) Melting peak análisis

Campylobacter spp (16S rRNA)

Chicken rinses

Buoyant density separation

Lightcycler probe

139

Campylobacter spp (16S rRNA)

Chicken skin

SE (Bolton broth)

Taqman probe

140

C. jejuni (VS1 sequence)

Chicken rinses

Direct detection

Taqman probe

141

C. jejuni (gyrA)

Chicken samples

Buoyant density separation

Melting peak análisis

142

Thermotolerant campylobacters (16S rRNA) C. jejuni (hipO), C. coli (ceuE) C. jejuni (Cj 0414)

Chicken rinses

Buoyant density separation

Lightcycler probe

143

Chicken carcasses Chicken carcasses

Direct detection Direct detection

Taqman probe Taqman probe

144 145

C. jejuni and C. coli (cadF)

Chicken carcasses

Qualitative PCR Direct detection

Gel electrophoresis

146

C. jejuni and C. coli (intergenic flaA-flaB sequence) C. jejuni (putative lipoprotein)

Minced beef, chicken, pork Chicken rinses

Rosef broth

Gel electrophoresis

147

Buoyant density separation

Gel electrophoresis

148

C. jejuni, C. coli, C. lari (16S rRNA) Campylobacter spp (16S rRNA)

Chicken skin, whole milk

Immunomagnetic separation

Biotin-avidin capture assay

149

Brocccoli, crabmeat,mushroom, raw milk, and raw oyster rinses Chicken samples

SE (blood-free enrichment broth)

Gel electrophoresis

150

SE (Preston broth)

Gel electrophoresis

151

Chicken samples

SE (Preston broth)

Gel electrophoresis

152

Raw meat and offal (poultry, porcine, ovine, and bovine), raw shellfish, milk Carcass rinse

SE (Campylobacter Enrichment Broth)

PCR ELISA DIG detection kit and two biotinylated capture probes

153

Campylobacter spp (16S rRNA), C. jejuni (mapA), C. coli (ceuE) Campylobacter spp (16S rRNA) C. jejuni and C. coli (ccoN)

C. jejuni and C. coli (ceuE)

Thermotolerant campylobacters Chicken carcass (16S rRNA) Thermophilic campylobacters Chicken legs, ground (23S rRNA), and C. jejuni (ceuE) beef, fermented sausage, roast beef, pork chops, turkey breast, chicken wieners, beef wieners Campylobacter spp (16S rRNA); Poultry samples C. jejuni (mapA), C. coli (ceuE) C. jejuni, C. coli (lpxA) Chicken rinses

138

Direct detection

PCR ELISA DIG detection kit

154

SE (Bolton broth)

Gel electrophoresis

155

SE (Bolton broth)

Gel electrophoresis

156

Direct detection and SE (Preston broth)

Gel electrophoresis

131

SE (Oxoid anaerobe basal broth)

Gel electrophoresis

157 (Continued)

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Molecular Detection of Foodborne Pathogens

Table 25.1  (Continued) Target

Matrix

Isolation Procedure

PCR Product Detection

Reference

C. jejuni, C. coli

Chicken rinses

DNA Array SE (Oxoid anaerobe basal broth)

Oligonucleotide probes (70mer)

157

Campylobacter spp (16S rRNA)

Chicken product

DNA FISH SE (Preston broth)

Oligonucleotide probe

152

Campylobacter spp (16S rRNA), thermophilic campylobacters (23S rRNA)

Chicken

SE (Preston broth)

Oligonucleotide probe

158

C. jejuni, C. coli, C. lari (16S rRNA)

Poultry products, dairy products, red meats, vegetables Chicken skin

NASBA SE (Preston broth)

ELGA

159

Direct detection and SE (Preston broth)

ELGA

160

Buoyant density separation

ELGA

161

Direct detection

Molecular beacon

162

C. jejuni, C. coli, C. lari (16S rRNA) C. jejuni (16S rRNA)

C. jejuni, C. coli (16S rRNA)

Poultry products, meat products, dairy products, water Poultry breast

Note:  SE, selective enrichment; ELGA, enzyme-linked gel assay.

T7 RNA polymerase. In contrast to PCR, the enzymes in NASBA work under isothermal conditions (41ºC) so a thermocycler is not necessary. The molecular target is singlestranded RNA and both ribosomal and messenger RNA may be amplified by NASBA. The amplification product is also single-stranded RNA but antisense to the original target. NASBA offers several advantages over PCR and RT-PCR. The number of copies generated by NASBA is higher than that of PCR and in a shorter time than RT-PCR. Cycles in NASBA exponentially increase the number of RNA copies so the sensibility is highest. Of special interest is that the background DNA does not interfere with the NASBA reaction. The likelihood of amplification of the contaminating DNA is prevented by the low temperature as NASBA reactions cannot exceed 41ºC without risk of enzyme denaturalization.167 Some disadvantages nevertheless are intrinsic to the NASBA reaction. It is not possible to control the extent of the reaction by adjusting the number of cycles because NASBA is isothermal. As a consequence the likelihood of nonspecific reactions increases and the product reaction can contain RNA fragments other than the specific copies of the target RNA. This creates the need of post-NASBA handling steps which increases both the time to identify the specific RNA product and the likelihood of molecular contamination. The product of NASBA reaction may be detected by gel electrophoresis followed by ethidium bromide staining, but a confirmatory step is needed to ensure product specificity. Enzyme-linked gel assay (ELGA) has been reported as a suitable method to identify the NASBA products by using a specific oligonucleotide probe (ELGA probe) 5′-end labeled with horseradish peroxidise.159,160 With the development of the real-time NASBA reactions this limitation was overcome.162,168 Real-time NASBA is based on beacon probes

which are small, single-stranded nucleic acids, ­hairpin probes that brightly fluoresce when they are bound to their targets.169 When NASBA is performed in combination with molecular probes, the target RNA products are detected in a sealed reaction tube, which simplifies analysis and eliminates an important source of assay contamination. Real-time NASBA has been used to detect C. jejuni in food samples with a high specificity and sensibility.162,168 In addition it was suitable to establish the viability of the molecular targets as 16S rRNA genes.162 (iii) Whole-genome-based methods: The whole-genomebased methods have been developed from the microarray technology that offers the possibility of depositing a high density of different specific probes on a very small area. They are also known as biochip, DNA chip, DNA microarray, and gene array. The DNA microarray consists of a series of samples spotted on a solid support, most commonly glass or silicon. Each spot contains DNA fragments or oligonucleotides as probes.170,171 Probes appear as spots in the final image where each spot represents a unique probe sequence and spots are usually 100–200 µm in size and located within 200–500 µm of each other. The targets are labelled with fluorescent dyes before to be applied to the microarray and the probe-target hybridization can be detected and quantified by fluorescence-based detection.172 Depending on the objective, targets may be PCR products, genomic DNA, total RNA, mRNA, cDNA, plasmid DNA, or oligonucleotides.173 The uses of the DNA arrays are likely unlimited as it can combine the amplification of nucleic acids with its massive screening capability. Simultaneous detection or level of expression of thousands of specific DNA can be achieved in a single DNA microarray, resulting in sensitivity, specificity, and highthroughput capacity.

353

Campylobacter

Two main types of arrays are produced: genomic arrays and oligonucleotide arrays. The genomic microarrays comprise either whole genome, genomic fragments from a cDNA library, or open reading frames from a bacterial strain. These arrays are useful for comparative phylogenetic studies and genetic diversity by determining constant and variable genes among the isolates. The oligonucleotide arrays, containing probes from 18 to 70 nucleotides long, 174 are useful for genomic analysis because these arrays can consist of multiple sequence variants of a target gene and can be adapted for pathogen detection in different environments. Although efforts to apply microarrays in food safety are important for detecting foodborne pathogens such as E. coli, Listeria, and others,175–177 Campylobacter detection in foods has not been well documented.178 To the best of our knowledge only one approach has been published157 that was suitable for specific identification of Arcobacter bultzleri, C. coli

Sample Preparation and DNA Extraction Preston Campylobacter Selective Enrichment Broth (Oxoid), containing Nutrient Broth No. 2, Preston Campylobacter Selective Supplement SR0117, Campylobacter Growth Supplement SR0232 and 5% (v/v) Lysed Horse Blood SR0048 Sterile plastic bags Orbital shaker 1.5 ml microcentrifuge tubes Micropipets and filter tips NucleoSpin Tissue kit (Macheray Nagel, Germany) 100% Ethanol ND-1000 Spectrophotometer (NanoDrop)

package liquid from whole chicken carcasses was labeled and hybridized to the microarray with and without enrichment. C. jejuni was detected efficiently both in package liquid from whole chicken carcasses and in enrichment broths.

25.2  Methods First, we describe a method for extracting DNA from Campylobacter spp. for PCR analysis. This method is suitable for extracting DNA directly from food samples. Extracting DNA with a commercial kit reduces the probability of contamination of the DNA with PCR inhibitors. Second, two PCR-based methods for detection of C. jejuni and C. coli are described: (i) multiplex PCR, and (ii) real-time duplex PCR.

25.2.1  Reagents and Equipment

PCR Amplification 5 U/µl Taq DNA polymerase (Invitrogen). The enzyme is supplied with 50 mM MgCl2 and 10 × reaction buffer (200 mM Tris pH 8.4, 500 mM KCl

100 mM stocks of each deoxyribonucleotide triphosphate (dNTPs; Bioline). dNTPS working stock solutions containing 10 mM of each dNTP The oligonucleotide primers and probes are listed in Table 25.2 Sterile ultrapure water 0.2 ml PCR tubes Thermal cycler Robocycler gradient 96 (Stratagene) Molecular grade agarose 10 × TBE buffer: 0.9 M Tris-HCl pH 8.3, 0.9 M boric acid, 0.02 M EDTA Agarose gel sample loading buffer: 40% sucrose, 0.25% bromophenol blue Agarose gel staining solution: 0.5 µg/ml (w/v) ethidium bromide in distilled water HyperLadder IV DNA molecular weight marker (Bioline) Standard apparatus for horizontal electrophoresis of agarose gels and power supply Gel documentation system ChemiDoc (BIO-RAD) Real-time thermocycler ABI PRISM 7000 Sequence Detection System (Applied Biosystems) MicroAmp optical tubes and optical caps (Applied Biosystems) 2 × Taqman Universal Master Mix kit (Applied Biosystems), containing AmpliTaq Gold DNA polymerase, dNTPs, Passive reference 1 (ROX), and optimized buffer components

and C. jejuni present in retail chicken samples using a DNA microarray without prior PCR as well as C. jejuni genotyping based on LOS class. The approach contains a comprehensive set of 70-mer oligonucleotide probes targeting genes implicated in metabolism or pathogenicity of those pathogens. Total genomic DNA isolated from the microbiota in the

25.2.2  Sample Preparation and DNA Extraction DNA is extracted using a commercial kit and follows the manufacturer’s instructions. The NucleoSpin Tissue kit (Macheray Nagel, Germany) is proposed as a method that provides a high grade of purity, and is suitable for

354

Molecular Detection of Foodborne Pathogens

PCR-based methods in Campylobacter detection from food samples. (1) Rinse 25 g of food sample in 100 ml of Preston broth and mix in an orbital shaker for 10 min at 200 rpm. (2) Centrifuge 1 ml culture for 10 min at 6,000 × g and discard the supernatant. (3) Resuspend the pellet in 180 μl Buffer T1. Add 25 μl proteinase K solution. Vortex vigorously and incubate at 56ºC in a shaking incubator until complete lysis is obtained (at least 30 min). (4) Add 200 μl Buffer B3, vortex vigorously and incubate at 70ºC for 10 min. (5) Add 210 μl 100% ethanol to the sample and vortex vigorously. (6) Place one NucleoSpin® Tissue Column into a collection tube. Apply the sample to the column. Centrifuge for 1 min at 11,000 × g. Discard the flow-through and place the column back into the Collection tube. (7) Add 500 μl Buffer BW. Centrifuge for 1 min at 11,000 × g. Discard flow-through and place the column back into the collection tube. (8) Add 600 μl Buffer B5 to the column and centrifuge for 1 min at 11,000 × g. Discard flow-through and place the column back into the collection tube. (9) Centrifuge the column for 1 min at 11,000 × g. (10) Place the NucleoSpin® Tissue Column into a 1.5-ml microcentrifuge tube and add 150 μl prewarmed Elution Buffer BE (70°C). Incubate at 70ºC for 1 min. Centrifuge 1 min at 11,000 × g.

(11) Determine DNA concentration in each sample using absorbance spectrophotometry. (12) Take an aliquot of the DNA suspension and dilute to give a stock template DNA concentration of 20 ng/µl. DNA suspensions may be stored at 4ºC or –20ºC.

25.2.3  Detection Procedures 25.2.3.1  Multiplex PCR The PCR conditions are those proposed by Denis et al.164 with minor modifications concerning the dNTPs concentration and 16S rRNA gene primer concentration.131 Multiplex PCR amplification for specific detection of the 16S rRNA (Campylobacter spp), mapA (C. jejuni), and ceuE (C. coli) genes is performed with the three primer sets described in Table 25.2. (1) Prepare PCR mixture (30-µl) containing 1 × PCR buffer, 1.5 mM MgCl2, 0.2 mM dNTPs, 0.6 U Taq polymerase, 0.5 µM of MD16S1 and MD16S2 primers, and 0.42 µM of MDmapA1, MDmapA2, COL3, and MDCOL2 primers (Table 25.3), and 100 ng of template DNA. (2) Run the PCR in a conventional thermal cycler using the following parameters: one cycle of 95ºC for 10 min; 35 cycles of 95ºC for 30 sec, 59ºC for 1 min 30 sec, 72ºC for 1 min; and one cycle of 72ºC for 10 min. (3) Mix 10 µl of the PCR product with 2 µl of gel loading buffer and put them into wells of 2% TBEagarose gel. In a parallel well, dispense 5 µl of the HyperLadder IV DNA molecular weight marker.

Table 25.2 Oligonucleotide Primers and Probes used for PCR Amplification and Detection Target

16S rRNA mapA ceuE

mapA

Name

MD16S1 MD16S2 MDmapA1 MDmapA2 COL3 MDCOL2

mapA-F mapA-R mapA-probe

ceuE

ceuE-F ceuE-R ceuE-probe

Sequence (5′→3′) Multiplex PCR ATCTAATGGCTTAACCATTAAAC GGACGGTAACTAGTTTAGTATT CTATTTTATTTTTGAGTGCTTGTG GCTTTATTTGCCATTTGTTTTATT AATTGAAAATTGCTCCAACTATG TGATTTTATTATTTGTAGCAGCG Duplex Real-Time PCR CTGGTGGTTTTGAAGCAAAGA CAATACCAGTGTCTAAAGTGCGTTTAT fam-TTGAATTCCAACATCGCTAATGTATAAAAGC CCTTT-tamra AAGCTCTTATTGTTCTAACCAATTCTAACA TCATCCACAGCATTGATTCCTAA vic-TTGGACCTCAATCTCGCTTTGGAATCATT-tamra

Note:  5′ fluorescent reporter dyes and 3′ quenchers underlined.

Size

Reference

857 bp

129

589 bp

179

462 bp

163

95 bp

180

102 bp

180

Campylobacter

(4) Subject to electrophoresis at 90 V for approx 1.5–2h in 1 × TBE buffer. (5) Remove gel and submerge in an ethidium bromide DNA staining solution (0.5 µg/ml) for 30 min. (6) Wash the gel with distilled water and visualize the DNA amplicons by exposure to a UV-transilluminator. (7) Expected product sizes are as follows: an 857-bp Campylobacter specific fragment of the 16S rRNA gene, and two species-specific fragments of 589 and 462-bp corresponding to C. jejuni and C. coli, respectively. 25.2.3.2  Duplex Real-Time PCR Duplex real-time PCR amplification for specific detection of the mapA (C. jejuni) and ceuE (C. coli) genes is performed with the two primer sets and Taqman probes described in Table 25.2. The protocol published by Best et al.180 involves a ceuE probe quantity higher (0.1 µM) than that employed by us (0.05 µM). Reducing the probe quantity to 0.05 µM significantly increases the specificity of the PCR. (1) Prepare PCR mixture (20-µl) containing 1 × Taqman Universal Master Mix, 0.3 µM of each primer, and 0.1 µM of mapA probe, 0.05 µM of ceuE probe (Table 25.2) and 100 ng of template DNA. (2) Add the PCR premix and DNA template to the MicroAmp optical tubes, and cap the tubes. (3) Perform PCR amplification in the thermocycler ABI PRISM 7000 with an initial denaturation/ enzyme activation at 95ºC for 10 min, followed by 40 cycles of two-step PCR consisting of 15-sec denaturation at 95ºC and 60-sec annealing/extension at 60ºC. (4) As with traditional PCR methods, include positive and negative controls for each pair of primers used. (5) After the run, detect PCR products directly by the Taqman machine monitoring the increase in fluorescence where a numerical value, the CT value (threshold cycle), is assigned. This indicates the cycle number at which measured fluorescence increased above calculated background fluorescence, identifying amplification of the target sequence.

25.3 Conclusion and Future Perspectives C. jejuni is now recognized as an important foodborne pathogen which is responsible of most bacterial gastroenteritis worldwide; its incidence is higher than those of infections caused by Salmonella species, Shigella species, or E. coli O157:H7. Despite its global importance insights into C. jejuni are limited compared to Salmonella and E. coli. As a consequence much effort has been directed toward an improved understanding of C. jejuni, its epidemiology and pathogenic mechanisms; however, many questions are yet to be answered.

355

C. jejuni is typically microaerophilic and moderately capnophilic which largely reduces the risk of growth in food products. However, human infections are usually acquired through the consumption of undercooked contaminated meat, water or other cross-contaminated food products, the foods of poultry origin being the most common sources of infections. How these organisms in both the environment and the food chain survive is an unanswered question. As they can enter a VBNC state under adverse conditions it has been suggested that this VBNC state can be a survival strategy in environments not supporting growth of Campylobacter. However, the significance of the VBNC-forms in transmission of disease is today uncertain, as the resuscitation back to the actively growing state is still controversial. Human infection, particularly in children, can be severe and normally associated with bloody diarrhea. Postinfectious complications are rare but debilitating long-term sequelae are associated with C. jejuni infections. Unlike most other foodborne pathogens C. jejuni is far less associated with general outbreaks. A variety of virulence factors have been identified, among them LOS structures resembling human neuronal gangliosides and CDT. But there is general agreement that C.  jejuni is unique in not having an identifiable enterotoxin as a mechanisms to cause diarrhea. The mechanism that causes diarrhea remain unclear. This limited understanding on C. jejuni pathogenesis is partly because of the lack of a suitable small animal model of infection that closely mimics human disease. Despite the progress that has been made in recent years, further research is required to provide the clues of how this pathogen survives in the environment and food as well as how it interacts with human cells. As exogenous microbiota C. jejuni can occur in small numbers in food and even with improved culture methods it could be very difficult to detect amid large numbers of indigenous microbiota in a complex sample matrix like food. Therefore, C. jejuni detection in foods by traditional culture methods, pre-enrichment and enrichment periods in selective media are absolutely neccessary. Further selective agar plating is also needed for colony isolation as well as species identification. In addition, there exist some VBNC-forms that are unable to grow on culture media. All of these contribute to the laborious and time-consuming features of the traditional culture ­methods as well as their inability to elucidate the VBNC-forms. Alternative rapid diagnoses including a wide range of novel testing procedures can significantly reduce the reporting time compared with conventional bacterial culture. In particular, methods targeting the nucleic acids, both DNA and RNA, are intrinsically more precise and less affected by natural variation than the phenotypic methods. In the past two decades, large numbers of both PCR-based methods (DNA amplification) and NASBA methods (RNA amplification) have been developed for C. jejuni detection; but the range of the applications of these methods for direct detection of C. jejuni in foods is narrow. The advent of whole-genome based methods, such as DNA microarrays, has generated new opportunities to detect pathogens in food.

356

Molecular Detection of Foodborne Pathogens

Although rapid, sensitive, and accurate the molecular methods also suffer from some disadvantages mainly when they are applied to the detection of pathogens in food. The complex matrix of the foods may interfere with both isolation and amplification of the target nucleic acid. The presence of inhibitory substances in the food products can affect primer binding or enzyme activity then resulting in false negative assays. Isolation of the DNA and RNA by in-house methods or commercial kits minimizes this limitation. The selection of the target to be amplified is another critical aspect when designing a molecular method. The target must be chosen on the basis of well established biological and genetic characteristics of the bacterium; data bases in the public domain from whole-genome sequencing are useful in that regard. However, a major disadvantage concerning molecular based methods is the question whether or not they are detecting nucleic acid from dead cells. As a result, validation, and optimization of in-house assays need further attention in the future. The uses of the DNA arrays are likely unlimited as it can combine the amplification of nucleic acids with its massive screening capability. Extending future research to this area should allow the identification of improved epidemiological markers which in turn should allow a rapid increase in the knowledge of the ecology, epidemiology, and pathogenesis of C. jejuni as well as future tools to reduce—if not eliminate—this important pathogen from the food chain.



Acknowledgments



Work in the authors’ laboratory is supported by grants from Spanish Government (AGL2005-08162-C02-01) and Basque Government (GIC07/52-IT-308-07).











References





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Campylobacter 112. Hald, B.K. et al. Study of the infectivity of saline-stored Campylobacter jejuni for day-old chicks. Appl. Environ. Microbiol., 67, 2388, 2001. 113. Ziprin, R.L. and Harvey, R.B. Inability of cecal microflora to promote reversion of viable non-culturable Campylobacter jejuni. Avian Dis. 48, 647, 2004. 114. Abubakar, I. et al. A systematic review of the clinical, public health and cost-effectiveness of rapid diagnostic tests for the detection and identification of bacterial intestinal pathogens in faeces and food. Health Technol. Assess., 11 (36), 2007. 115. Tolcin, R. et al. Evaluation of the Alexon-trend ProSpecT Campylobacter microplate assay. J. Clin. Microbiol., 38, 3853, 2000. 116. Dediste. A. et al. Evaluation of the ProSpecT microplate assay for detection of Campylobacter: a routine laboratory perspective. Clin. Microbiol. Infect., 9, 1085, 2003. 117. Mouwen, D.J.M. et al. Discrimination of enterobacterial repetitive intergenic consensus PCR types of Campylobacter coli and Campylobacter jejuni by Fourier Transform Infrared Spectroscopy. Appl. Environ. Microbiol., 71, 4318, 2005. 118. Mouwen, D.J.M. et al. Artificial neural network based identification of Campylobacter species by Fourier transform infrared spectroscopy. J. Microbiol. Methods, 67, 131, 2006. 119. Cook, N, The use of NASBA for the detection of microbial pathogens in food and environmental samples. J. Microbiol. Methods, 53, 165, 2003. 120. Lubeck, P.S. et al. Toward an international standard for PCRbased detection of food-borne thermotolerant Campylobacters: assay development and analytical validation. Appl. Environ. Microbiol., 69, 5664, 2003. 121. Alonso, R. et al. PCR-restriction fragment length polymorphism assay for detection of gyra mutations associated with fluoroquinolone resistance in Campylobacter coli. Antimicrob. Agents Chemother., 48, 4886, 2004. 122. Amar, C.F. et al. Blinded application of microscopy, bacteriological culture, immunoassays and PCR to detect gastrointestinal pathogens from faecal samples of patients with community-acquired diarrhoea. Eur. J. Clin. Microbiol. Infect. Dis., 23, 529, 2004. 123. Alonso, R. et al. MAMA-PCR assay for the detection of point mutations associated with high-level erythromycin resistance in Campylobacter jejuni and Campylobacter coli strains. J. Microbiol. Methods, 63, 99, 2005. 124. Jensen, A.N. et al. Development of real-time PCR and hybridization methods for detection and identification of thermophilic Campylobacter spp. in pig faecal samples. J. Appl. Microbiol., 99, 292, 2005. 125. Kostic, T. et al. A microbial diagnostic microarray technique for the sensitive detection and identification of pathogenic bacteria in a background of nonpathogens. Anal. Biochem., 360, 244, 2007. 126. O’Sullivan, N.A. et al. Detection and differentiation of Campylobacter jejuni and Campylobacter coli in broiler chicken samples using a PCR/DNA probe membrane based colorimetric detection assay. Mol. Cell. Probes, 14, 7, 2000. 127. Nayak, R., Stewart, T.M., and Nawaz, M.S. PCR identification of Campylobacter coli and Campylobacter jejuni by partial sequencing of virulence genes. Mol. Cell. Probes, 19, 187, 2005. 128. Heid, C.A. et al. Real time quantitative PCR. Genome Res., 6, 986, 1996.

359 129. Linton, D. et al. PCR detection, identification to ­species level, and fingerprinting of Campylobacter jejuni and Campylobacter coli direct from diarrheic samples. J. Clin. Microbiol., 35, 2568, 1997. 130. On, S.L., and Jordan, P.J. Evaluation of 11 PCR assays for species-level identification of Campylobacter jejuni and Campylobacter coli. J. Clin. Microbiol., 41, 330, 2003. 131. Mateo, E. et al. Evaluation of a PCR assay for the detection and identification of Campylobacter jejuni and Campylobacter coli in retail poultry products. Res. Microbiol., 156, 568, 2005. 132. Martinez, I. et al. Detection of cdtA, cdtB, and cdtC genes in Campylobacter jejuni by multiplex PCR. Int. J. Med. Microbiol., 296, 45, 2006. 133. Cheng, Z. and Griffiths, M.W. Rapid detection of Campylobacter jejuni in chicken rinse water by melting-peak analysis of amplicons in real-time polymerase chain reaction. J. Food Prot., 66, 1343, 2003. 134. Sails, A.D. et al. A real-time PCR assay for the detection of Campylobacter jejuni in foods after enrichment culture. Appl. Environ. Microbiol., 69, 1383, 2003. 135. Josefsen, M.H., Jacobsen, N.R. and Hoorfar, J. Enrichment followed by quantitative PCR both for rapid detection and as a tool for quantitative risk assessment of food-borne thermotolerant campylobacters. Appl. Environ. Microbiol., 70, 3588, 2004. 136. Oliveira, T.C., Barbut, S. and Griffiths, M.W. Detection of Campylobacter jejuni in naturally contaminated chicken skin by melting peak analysis of amplicons in real-time PCR. Int. J. Food Microbiol., 104, 105, 2005. 137. Abu-Halaweh, M., Bates, J. and Patel, B.K. Rapid detection and differentiation of pathogenic Campylobacter jejuni and Campylobacter coli by real-time PCR. Res. Microbiol., 156, 107, 2005. 138. Oliveira, T.C., Barbut, S. and Griffiths, M.W. A robotic DNA purification protocol and real-time PCR for the detection of Campylobacter jejuni in foods. J. Food Prot., 68, 2131, 2005. 139. Wolffs, P. et al. Quantification of Campylobacter spp. in chicken rinse samples by using flotation prior to real-time PCR. Appl. Environ. Microbiol., 71, 5759, 2005. 140. Krause, M. et al. Comparative, collaborative, and on-site validation of a TaqMan PCR method as a tool for certified production of fresh, campylobacter-free chickens. Appl. Environ. Microbiol., 72, 5463, 2006. 141. Debretsion, A. et al. Real-time PCR assay for rapid detection and quantification of Campylobacter jejuni on chicken rinses from poultry processing plant. Mol. Cell Probes, 21, 177, 2007. 142. Fukushima, H. et al. Rapid separation and concentration of food-borne pathogens in food samples prior to quantification by viable-cell counting and real-time PCR. Appl. Environ. Microbiol., 73, 92, 2007. 143. Wolffs, P.F.G. et al. Simultaneous quantification of pathogenic Campylobacter and Salmonella in chicken rinse fluid by a flotation and real-time multiplex PCR procedure. Int. J. Food Microbiol., 117, 50, 2007. 144. Hong, J. et al. Quantification and differentiation of Campylobacter jejuni and Campylobacter coli in raw chicken meats using a real-time PCR method. J. Food Prot., 70, 2015, 2007. 145. Ronner, A.C. and Lindmark, H. Quantitative detection of Campylobacter jejuni on fresh chicken carcasses by real-time PCR. J. Food Prot., 70, 1373, 2007.

360 146. Konkel, M.E. et al. Identification of the enteropathogens Campylobacter jejuni and Campylobacter coli based on the cadF virulence gene and its product. J. Clin. Microbiol., 37, 510, 1999. 147. Waage, A.S. et al. Detection of small numbers of Campylobacter jejuni and Campylobacter coli cells in environmental water, sewage, and food samples by a seminested PCR assay. Appl. Environ. Microbiol., 65, 1636, 1999. 148. Wang, H. et al. Improved PCR detection of Campylobacter jejuni from chicken rinses by a simple sample preparation procedure. Int. J. Food Microbiol., 52, 39, 1999. 149. Waller, D.F. and Ogata, S.A. Quantitative immunocapture PCR assay for detection of Campylobacter jejuni in foods. Appl. Environ. Microbiol., 66, 4115, 2000. 150. Thunberg, R.L., Tran, T.T. and Walderhaug, M.O. Detection of thermophilic Campylobacter spp. in blood-free enriched samples of inoculated foods by the polymerase chain reaction. J. Food Prot., 63, 299, 2000. 151. Denis, M. et al. Campylobacter contamination in French chicken production from farm to consumers. Use of a PCR assay for detection and identification of Campylobacter jejuni and Campylobacter coli. J. Appl. Microbiol., 91, 255, 2001. 152. Moreno, Y. et al. Direct detection of thermotolerant campylobacters in chicken products by PCR and in situ hybridization. Res. Microbiol., 152, 577, 2001. 153. Bolton, F.J. et al. Detection of Campylobacter jejuni and Campylobacter coli in foods by enrichment culture and polymerase chain reaction enzyme-linked immunosorbent assay. J. Food Prot., 65, 760, 2002. 154. Hong, Y. et al. Rapid detection of Campylobacter coli, C. jejuni, and Salmonella enterica on poultry carcasses by using PCR-enzyme-linked immunosorbent assay, Appl. Environ. Microbiol., 69, 3492, 2003. 155. Josefsen, M.H. et al. Validation of a PCR-based method for detection of food-borne thermotolerant campylobacters in a multicenter collaborative trial. Appl. Environ. Microbiol., 70, 4379, 2004. 156. Bohaychuk, V.M. et al. Evaluation of detection methods for screening meat and poultry products for the presence of foodborne pathogens. J. Food Prot., 68, 2637, 2005. 157. Quinones, B. et al. Detection and genotyping of Arcobacter and Campylobacter isolates from retail chicken samples by use of DNA oligonucleotide arrays. Appl. Environ. Microbiol., 73, 3645, 2007. 158. Schmid, M.W. et al. Development and application of oligonucleotide probes for in situ detection of thermotolerant Campylobacter in chicken faecal and liver samples. Int. J. Food Microbiol., 105, 245, 2005. 159. Uyttendaele, M. et al. Detection of Campylobacter jejuni added to foods by using a combined selective enrichment and nucleic acid sequence-based amplification (NASBA). Appl. Environ. Microbiol., 61, 1341, 1995. 160. Uyttendaele, M., Bastiaansen, A. and Debevere, J. Evaluation of the NASBA nucleic acid amplification system for assessment of the viability of Campylobacter jejuni. Int. J. Food Microbiol., 37, 13, 1997. 161. Uyttendaele, M., Debevere, J. and Lindqvist, R. Evaluation of buoyant density centrifugation as a sample preparation method for NASBA–ELGA detection of Campylobacter jejuni in foods. Food Microbiol., 16, 575, 1999.

Molecular Detection of Foodborne Pathogens 162. Churruca, E. et al. Detection of Campylobacter jejuni and Campylobacter coli in chicken meat samples by real-time nucleic acid sequence-based amplification with molecular beacons. Int. J. Food Microbiol., 117, 85, 2007. 163. Gonzalez, I. et al. Specific identification of the enteropathogens Campylobacter jejuni and Campylobacter coli using a PCR test based on the ceuE gene encoding a putative virulence determinant. J. Clin. Microbiol., 35, 759, 1997. 164. Denis, M. et al. Development of a m-PCR assay for simultaneous identification of Campylobacter jejuni and C. coli. Lett. Appl. Microbiol., 29, 406, 1999. 165. Keer, J.T., and Birch, L. Molecular methods for the assessment of bacterial viability. J. Microbiol. Methods, 53, 175, 2003. 166. Kievits, T. et al.,. NASBA isothermal enzymatic in vitro nucleic-acid amplification optimized for the diagnosis of HIV-1 infection. J. Virol. Methods, 35, 273, 1991. 167. Tai, J.H. et al. Development of a rapid method using nucleic acid sequence-based amplification for the detection of astrovirus. J. Virol. Methods, 110, 119, 2003. 168. Cools, I. et al. Development of a real-time NASBA assay for the detection of Campylobacter jejuni cells. J. Microbiol. Methods, 66, 313, 2006. 169. Tyagi, S. and Kramer, F.R. Molecular beacons: probes that fluoresce upon hybridization. Nat. Biotechnol., 14, 303, 1996. 170. Al-Khaldi, S.F. et al. DNA microarray technology used for studying foodborne pathogens and microbial habitats: minireview. J. AOAC Int., 85, 906, 2002. 171. Ramsay, G. DNA chips: state-of-the art. Nat. Biotechnol., 16, 40, 1998. 172. Vora, G. J. et al. Nucleic acid amplification strategies for DNA microarray-based pathogen detection. Appl. Environ Microbiol., 70, 3047, 2004. 173. Call, D.R., Borucki, M.K. and Loge, F.J. Detection of bacterial pathogens in environmental samples using DNA microarrays. J. Microbiol. Methods, 53, 235, 2003. 174. Bryant, P.A. et al. Chips with everything: DNA microarrays in infectious diseases. Lancet, 4, 100, 2004. 175. Delaquis, P. et al. Application of DNA microarrays for the identification and tracking of E. coli O157:H7 and Listeria monocytogenes in food production systems. In Meeting of Agriculture and Agri-Food Canada Food Network, Lacombe, Alberta, 2002, p. 23 [Abstr. 8]. 176. Wilson, W.J. et al. Sequence-specific identification of 18 pathogenic microorganisms using microarray technology. Mol. Cell. Probes, 16, 119, 2002. 177. Hong, B. et al. Application of oligonucleotide array technology for the rapid detection of pathogenic bacteria of foodborne infections. J. Microbiol. Methods, 58, 403, 2004. 178. Kostrzynska, M. and Bachand, A. Application of DNA microarray technology for detection, identification, and characterization of food-borne pathogens. Can. J. Microbiol., 52, 1, 2006. 179. Stucki, U. et al. Identification of Campylobacter jejuni on the basis of a species-specific gene that encodes a membrane protein. J. Clin. Microbiol., 33, 855, 1995. 180. Best E.L. et al. Applicability of a rapid duplex real-time PCR assay for speciation of Campylobacter jejuni and Campylobacter coli directly from culture plates. FEMS Microbiol. Lett., 229, 237, 2003.

26 Enterobacter

Angelika Lehner and Roger Stephan University of Zurich

Carol Iversen and Seamus Fanning University College Dublin

Contents 26.1 Introduction.................................................................................................................................................................... 361 26.1.1 Classification..................................................................................................................................................... 361 26.1.2 Epidemiology and Pathogenesis....................................................................................................................... 361 26.1.3 Diagnosis........................................................................................................................................................... 362 26.1.3.1 Conventional Procedures................................................................................................................. 362 26.1.3.2 Molecular Procedures...................................................................................................................... 363 26.2 Methods.......................................................................................................................................................................... 364 26.2.1 Sample Preparation........................................................................................................................................... 364 26.2.2 Detection Procedures........................................................................................................................................ 365 26.2.2.1 PCR Identification .......................................................................................................................... 365 26.2.2.2 Real-Time PCR Identification.......................................................................................................... 365 26.3 Conclusions and Future Perspectives............................................................................................................................. 365 References.................................................................................................................................................................................. 365

26.1  Introduction 26.1.1 Classification The Enterobacter genus represents a large and heterogeneous group within the Enterobacteriaceae family. It once had 21 taxonomically valid species, however recent taxonomic studies have led to the transfer of three Enterobacter species to alternative genera (E. intermedius=Kluyvera intermedia, E. agglomerans=Pantoea agglomerans, E. sakazakii=Cronobacter spp.). Additionally, E. taylorae has been recognized as a heterotypic synonym of E. cancerogenus and Enterobacter dissolvens has been reassigned to Enterobacter cloacae as E. cloacae subspecies dissolvens comb. nov. Thus, the genus Enterobacter currently contains 16 distinct species.1–3 Species closely related to E. cloacae, having a DNA relatedness of over 60% to E. cloacae and/or differing by only one biochemical trait have been subsumed in the so-called E. cloacae-complex. These are considered the main species that are frequently isolated from clinical samples.4 Members of the genus Enterobacter can be found in natural environments (soil, water, sewage, vegetables) and sometimes appear as commensals of the animal and human gut. Only one species—Enterobacter sakazakii—is recognized as a foodborne pathogen and therefore we will focus in this chapter mainly on this species. E. sakazakii, previously known as “yellow-pigmented Enterobacter cloacae,” was

described as a new species in 1980 by Farmer et al.5 when the presence of certain biochemical traits, antibiotic susceptibilities and DNA relatedness provided sufficient evidence to distinguish the species from E. cloacae. However, from the beginning, members of this species were described as quite heterogeneous at the microbiological as well as at the molecular level.5–7 Updating the original taxonomy of E. sakazakii by using a polyphasic approach has resulted in the definition of at least five new species based on extensive geno- and phenotypic evaluations.8,9 In order to facilitate their continued inclusion in schemata for the diagnosis and the microbiological criteria for foodstuffs, it has been proposed that these species be moved to a novel genus, Cronobacter 9,10 with the novel genus being synonymous with E. sakazakii. A further study based on multilocus sequence analysis (MLSA) of several house keeping genes present in E. sakazakii supports the classification of these organisms as a new genus (Kuhnert et al., in submission).

26.1.2 Epidemiology and Pathogenesis Although isolated with varying frequency from the natural, domestic, and food production environments, as well as from various types of food, E. sakazakii attracted special attention as an (occasional) contaminant of powdered infant formula (PIF).11–13 Multiple cases of infections have occurred in neonatal intensive care units (NICU) and a number of outbreaks 361

362

have been traced back to the presence of these pathogens in reconstituted infant formula milk and/or their persistence in/ on food preparation equipment.14–16 A summary of reported cases has been provided by Mullane et al.17 Enterobacter sakazakii can cause meningitis, necrotizing enterocolitis (NEC) and bacteremia especially in neonates and infants.16,18–20 The International Commission for Microbiological Specification for Foods (ICMSF, 2002) has ranked E. sakazakii as “Severe hazard for restricted populations, life threatening or substantial chronic sequelae or long duration.” Although it is possible for infection to be acquired by previously healthy newborn infants in the home environment,21,22 the majority of reported cases have occurred in NICU. A retrospective study of 46 infants indicated that meningitis is more prevalent in infants of normal gestational age and birth weight with onset of disease usually occurring within the first week following birth. In contrast low birth weight, premature infants were more likely to develop bacteremia with no progression to CNS disease and the age of onset was usually over 1 month.23 A higher mortality rate and adverse sequelae in survivors were associated with meningitis cases. The bacterium causes cystic changes, abscesses, fluid collection, dilated ventricles and infarctions. Cronobacter meningitis leads to cerebral abscess similar to those due to Citrobacter koseri infections, therefore a similarity in the cascade of pathogenic events induced by the two organisms has been suggested.24 Some Cronobacter infections occur in neonates born by Cesarean section.18,25–27 Therefore, it is thought risk of infection is related to ingestion, exacerbated by poor hygiene practices, and not through vertical transmission from the mother.26 Prolonged and repeated use of enteral feed bags are identified as risk factors28,29 with the possibility of biofilm formation leading to increased oral dose. Muytjens et al.18 re-evaluated Enterobacter strains from blood and CSF and uncovered several cases of meningitis and bacteremia due to Cronobacter infection suggesting that the organism had been under reported. The first known cases of meningitis due to Cronobacter occurred in England.25 No source for the infection was identified although the report does not mention epidemiological investigation of infant formula or feed preparation equipment. Since then cases have been reported worldwide. Cronobacter has been isolated from a wide range of clinical sources including a stethoscope5 and nursery food preparation equipment.14,27,30 Smeets et al.31 used pulsed-field gel electrophoresis (PFGE) to confirm the epidemiological evidence that a contaminated dish brush used for cleaning bottles was the source of three cases in 1981. In Iceland three cases were reported linked to milk formula contaminated with Cronobacter 32 Two groups14,15 reported on four neonates with Cronobacter infections in Tennessee. The organism was isolated from all four patients, a used can of infant formula milk and the blender (which had heavy growth of the organism). In this outbreak identical biotypes, antibiograms, and plasmid profiles were obtained for patients and environmental isolates. There was evidence of prolonged incubation in bottle heaters between 35–37°C before use. Nazarowec-White and Farber33 studying three isolates

Molecular Detection of Foodborne Pathogens

obtained from one hospital over 11 years showed that they had indistinguishable ribotype patterns indicating possible persistence in the environment. In 1994 an outbreak occurred in France involving 17 neonates. Cronobacter isolates were obtained from various anatomical sites, prepared feeds and unused infant formula. PFGE analysis described four clusters, two of which contained isolates from neonates who ranged from asymptomatic colonized individuals to case fatalities. One other cluster contained isolates from the prepared feed, an asymptomatic neonate and a neonate with mild digestive problems. The fourth cluster comprised the isolates from the unused powdered formula. In this case no link could be made between the powdered formula and the outbreak and it is possible that the prepared feed became contaminated via an alternate source.34 A further outbreak occurred in France in 2004 involving 13 infants, nine were asymptomatically colonized, two developed bacterial meningitis, one had conjunctivitis and one suffered hemorrhagic colitis. The outbreak was investigated using automated ribotyping (using restriction endonucleases EcoRI, PstI, and PvuII) and by PFGE using the enzymes Xba1 and Spe1. A total of nine isolates from eight neonates were found to have undistinguishable ribotypes and PFGE patterns. These isolates were undistinguishable from isolates obtained from four separate lots of infant formula, thus establishing a clear link between the outbreak and the product.79 NEC is much more common in babies fed formula milk compared with those fed breast milk.35 The pathogenesis is associated with neonatal intestinal ischemia, microbial colonization of the gut, and excess protein substrate in the intestinal lumen (the latter being associated with formula feeding). In a study of 125 neonates with NEC, Enterobacter spp. were the most common organisms, being isolated from 29% of patients.36 As with cases of meningitis, NEC due to Cronobacter traditionally has a high case fatality rate (10–55%). Van Acker et al.16 described 12 cases of NEC in neonates that occurred in 1998. Eleven strains of Cronobacter were isolated from patient samples and 14 strains were isolated from infant milk preparations. Arbitrary primed PCR (AP-PCR) was used to type all the Cronobacter isolates and determine common sources of the outbreak. Three AP-PCR profiles were obtained for patient and milk isolates with the 14 milk isolates matching the profile from three patients. Four years earlier E. sakazakii had been isolated from a gastrostomy tube of a neonate fed the same type of milk. This original isolate was subsequently shown to have an AP-PCR profile almost identical to the 14 milk and patient isolates, thus demonstrating a persistent contamination problem.

26.1.3  Diagnosis 26.1.3.1  Conventional Procedures In 2002, the Food and Drug Administration (FDA) published a method for detection of E. sakazakii which included a ­pre-enrichment step in buffered peptone water (BPW), enrichment in Enterobacteriaceae Enrichment (EE) broth, plating on Violet Red Bile Glucose agar (VRBG) and picking

Enterobacter

of five typical colonies onto tryptone soy agar (TSA) plates.37 After incubation at 25°C for 48–72 h, yellow pigmented colonies on TSA plates are confirmed using the API 20E system. The main weak points of this procedure are the inability of some target strains to grow in the selective EE broth, the lack of discrimination between Enterobacteriaceae strains on VRBG, and the variation in intensity of the pigmentation, with occasional observation of nonpigmented strains.5,10 Guillaume-Gentil et al.38 published an alternative procedure based on selective enrichment in a modified lauryl sulphate tryptose broth (mLST), incorporating 0.5 M NaCl, and 10 mg/ml vancomycin hydrochloride. This method was further improved by the replacement of VRBG, with a chromogenic agar and forms the basis of the ISO Technical Specification for detection of E. sakazakii in milk-based infant formula.13 Various chromogenic and fluorogenic agar media have been described in recent years for detection of E. sakazakii.39–43 These are based mainly on the enzyme α-glucosidase, which is constitutively expressed in E. sakazakii. However, other organisms also produce presumptive colonies on these agars, notably the newly described species E. helveticus, E. turicensis, and E. pulveris.2,3 These species can be found in the same ecological niches as E. sakazakii, such as dried food products and factory environments and present a challenge to both culture-based as well as molecular isolation and identification methods. It has been established that some isolates of E. sakazakii do not grow well in enrichment broths currently proposed for isolation of this organism.44,45 As some E. sakazakii strains cannot grow in these broths, a sample containing only such strains would give a false negative result and therefore the enrichment procedure should be improved. In a recent study a Cronobacter Screening Broth (CSB), has been developed to identify samples potentially contaminated with Cronobacter (E. sakazakii).46 The broth is designed to circumvent the problems encountered with selective enrichment media for these organisms and to be complementary to current available chromogenic media in order to improve overall sensitivity and selectivity of Cronobacter (E. sakazakii) detection. An alternative method to avoid the difficulties of selective broths is the MATRIX PSAK50 Method (Matrix MicroScience Ltd, UK), which uses cationic paramagnetic particle capture to concentrate contaminating microorganism in a pre-enriched sample before plating directly onto isolation agar.47 Currently the AOAC® Official Methods ProgramSM is in the process of assessing rapid methods for the detection of E. sakazakii. These include two enzyme-linked immunoassays (EIAs): the Assurance for Enterobacter sakazakii (BioControl Systems, USA) and the TECRA HELIX E. sakazakii Method (TECRA International, Australia). 26.1.3.2  Molecular Procedures (i) Species-specific detection and identification: In comparison with labor-intensive methods, the recently developed fluoro- and chromogenic, differential (selective) media decrease the time for isolation of E. sakazakii.39–42,44

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However, due to the obvious heterogeneity within these organisms and/or the fact that expression of phenotypical/ biochemical features may vary, the phenotypic identification of E. sakazakii with available commercial systems may be difficult.45 Molecular methods revealed that several strains identified as E. sakazakii by commercial biochemical kits belonged to distinct species.7,42,48 Restaino et al.42 recommended using more than one differentiation system. Several conventional and real-time PCR methods have been developed that enable (quantitative), sensitive, specific, and rapid detection from infant formula, enrichment broths and media. Targets for conventional PCR systems include the 16S rRNA gene,6,49,50 the ompA gene51 the gene coding for the 1,6 α-glucosidase in E. sakazakii52 and a gene encoding a zinc-containing metalloprotease.53 Of the “real-time”-assays that have been described so far, three of them target the 16S rRNA gene,54–56 one is based on the region located between the 16S rRNA and the 23 rRNA genes,57 another targets the region between the tRNA-glu and 23S rRNA genes58 and one targets the dnaG gene.48,59 To date, only two PCR based methods have been evaluated on a broad panel of target as well as nontarget strains and both of these proved to be 100% sensitive and specific for E. sakazakii. These methods will be described in detail in the methods section of this chapter. Genetic-based systems include the BAX® Assay for the Detection of Enterobacter sakazakii (Dupont Qualicon,USA) and the foodproof® Enterobacteriaceae plus E. sakazakii Detection System (BIOTECON Diagnostics, Germany). The latter qualitatively detects Enterobacteriaceae DNA while simultaneously identifying the presence of E. sakazakii. To avoid false-positive PCR results the DNA of dead bacterial cells in the sample is eliminated using a light sensitive reagent that only penetrates the cell membranes of dead cells and covalently binds to DNA preventing amplification. The US FDA have also developed a revised BAM Method involving a short enrichment step, centrifugation, and a PCR assay. Another commercially available PCRbased identification system is the TaqMan® Enterobacter sakazakii Detection Kit (Applied BioSystems, USA). The VIT® (vermicon identification technology, Munich, Germany) represents an alternative to the DNA-targeted PCRbased detection and identification systems for E. ­sakazakii. It is based on fluorescently labeled gene probes targeting specified regions on the ribosomal RNA of the bacteria, therefore only live cells are detected by the system. The test is performed on 1 ml of overnight culture of enrichment broth or rich media broth (e.g., BHI, LB) after inoculation with presumptive colony material according to the instructions of the manufacturer. The method has successfully been used in a comparative study evaluating cultural and molecular identification systems for E. sakazakii 44. (ii) Subtyping: The determination of clonality or polyclonality of isolates is necessary during investigations of E. sakazakii infection outbreaks and also for trace-back studies in production environments.

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PFGE. PFGE is a nonamplified technique that separates long strands of DNA molecules (larger than 15–20 kb) previously digested with restriction enzyme(s). Periodical switching of the voltage in three directions results in the reorientation of DNA moving through the gel in a size dependent manner, which facilitates a finer resolution as it aligns and realigns to the applied electrical field. Nazarowec-White and Farber (1999)33 were the first to describe a PFGE method for the subtyping of E. sakazakii isolates and used two different enzymes (XbaI and SpeI). Recently, two factory surveillance studies have employed a PFGE method for tracing E. sakazakii isolates within infant food manufacturing facilities (Iversen et al., in submission and Mullane et al.60). These studies were based on the protocol of Nazarowec-White and Farber33 and used XbaI. In 2008, a collaborative study was undertaken by several international laboratories, in conjunction with the PulseNet Programme at the Centres for Disease Control and Prevention (CDC) in the US, to develop a standard protocol for E. sakazakii PFGE typing. Ribotyping. Ribotypes are generated by probing restriction fragments of genomic DNA for the highly conserved genes coding for the 16S and 23S rRNA. Small variations between strains occur in the less conserved, flanking genes and intergenic sections of the genome resulting in fragments of unequal size. These are separated using gel electrophoresis, transferred to a membrane and the individual strain fingerprints revealed by hybridization of chemiluminescent probes. The automated RiboPrinter Microbial Characterization System (Dupont Qualicon, USA) has been used to speciate and characterize a large number of E. sakazakii isolates.8 Isolates were grown on TSA (18 h, 37°C) and prepared according to standard procedures61 using the EcoR1 restriction enzyme. Riboprint patterns are analyzed based on the number, size, and signal intensity of the detected fragments. Comparison to existing entries in a riboprint database allows species-and subspecies-level identification. Random amplification of polymorphic DNA (RAPD). RAPD typing has been used to analyze clonal relationships between E. sakazakii strains in a number of independent studies (Iversen et al., in submission).33,48,62–65 NazarowecWhite and Farber33 developed protocols for the molecular subtyping of E. sakazakii by ribotyping, RAPD, and PFGE. The authors showed that RAPD and PFGE were the most discriminatory subtyping schemes for E. sakazakii followed by ribotyping and two microbiological typing methods— biotyping and antibiograms. In addition, Clementino et al.62 comparatively assessed the usefulness of the tRNA intergenic spacer, 16S–23S internal transcribed spacer and randomly amplified DNA for discrimination of E. sakazakii and E. cloacae isolates. BOX-PCR and REP-PCR. Proudy et al.66 examined the discriminative power of the 154 bp BOX element against the sequencing of the fliC gene and PFGE using 27 E. sakazakii strains from clinical and environmental sources. The BOXPCR results showed 92% agreement with PFGE results indicating the potential of this typing method for ­epidemiological investigation, whereas fliC gene sequencing was poorly

Molecular Detection of Foodborne Pathogens

discriminative. The application of BOX-PCR genotyping in a factory environment was demonstrated in a follow-up study.67 More recently the discriminative power of the repPCR (repetitive extragenic palindromic (REP) has been compared to PFGE.68 Using Simpson’s index of diversity, values of 0.974 and 0.998 were calculated for rep-PCR and PFGE, respectively at a similarity cut-off of 95% demonstrating good correlation with a high degree of genetic heterogeneity among the isolates. Multi-locus VNTR analysis (MLVA). Variable number tandem repeat (VNTR) motifs represent sources of genetic polymorphisms. These DNA sequence elements are often maintained within a bacterial species, with individual strains displaying different copy numbers. The length of a tandem repeat at a specific locus can vary as a consequence of DNA slippage during replication or unequal crossover elements. These differences can be analyzed by amplification of the region and sizing of the resulting amplicons.69 The high degree of polymorphism at these loci is particularly useful as a target for strain discrimination within bacterial species. MLVA is a subtyping method that involves amplification and fragment size comparison of polymorphic VNTR regions and has successfully been used to type Enterobacteriaceae.70–72 The availability of a complete E. sakazakii genome sequence (http://genome/wustl.edu/pub/organism/Microbes/Enteric_ Bacteria/Enterobacter_sakazakii/assembly/Enterobacter_ sakazakii-4.0/) enabled the identification of VNTR motifs within E. sakazakii. Subsequently an MLVA subtyping scheme was developed and applied on a genotypically and phenotypically diverse collection of E. sakazakii isolates in a study by Mullane et al.73 Amplified fragment length polymorphisms (AFLP). The AFLP technique has been employed in plant and microbiological research to describe the molecular ecology of various niches and can be used to determine inter- and intraspecies relatedness.74–76 Mougel et al.77 found that members of the same genomic species cluster consistently using AFLP analysis and suggested that future genomic delineation of bacterial species could be based on this approach. This technique was included in a study by Iversen et al.8 to clarify the taxonomic relationship of over 200 strains previously identified as E. sakazakii and gave discriminatory results comparable to DNA–DNA hybridization.

26.2  Methods 26.2.1 Sample Preparation Aliquots of matrices to be tested are pre-enriched in 9 × volume of BPW for 24 h at 37°C. A 100-fold dilution (0.1 ml in 10 ml) is transferred into CSB (CSB: 18 g/l Bromocressol purple broth, Sigma B2676; 10 g/l sucrose and 10 mg/l vancomycin hydrochloride, Sigma V2002)46 or alternative enrichment broths as e.g., mLST.13 Fermentation of the carbohydrate results in a colour change from purple to yellow. After incubation at 41.5°C for up to 24 h, positive (yellow) broths are streaked onto chromogenic media,

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Enterobacter

e.g., Brilliance Enterobacter sakazakii agar (CM1055, Oxoid, UK), Enterobacter sakazakii Isolation Agar (ESIA, AES Cheminux, France), Enterobacter sakazakii Screening Plate (ChromID Sakazakii, bioMérieux, France), or equivalent. For DNA extraction from broth cultures, commercially available kits e.g., the Qiagen Blood and Tissue Kit (Qiagen, Germany) are recommended. PCR and real-time PCR assays are set up using standard molecular handling procedures and by applying appropriate equipment and using standard instruments.

26.2.2  Detection Procedures 26.2.2.1  PCR Identification In a study by Lehner et al.52 the molecular basis of the α-glucosidase activity in E. sakazakii was determined and the potential of this PCR system targeting the 1,6-αglucosidase for the specific identification of E. sakazakii was evaluated in two studies.10,44 Presumptive colonies from agar plates are inoculated into 5 ml BHI broth and grown overnight at 37°C. DNA is extracted using a simple “boiling method,” i.e., harvesting cells from 1 ml overnight culture by centrifugation (5  min at 7500 × g), resuspension of cells in 100 µl distilled water, lysis of cells by heating the suspension at 100°C for 10 min and final separation of DNA from cellular debris by centrifugation for 2 min at 10000 × g. Alternatively, DNA can be extracted using a commercially available kit (e.g., Qiagen Blood and Tissue Kit, Germany). The α-glucosidase (gluA) gene is amplified using the following primers: EsAgf: 5′-TGAAAGCAATCGACAAGAAG-3′ and EsAgr: 5′-ACTCATTACCCCTCCTGATG-3′ generating a product of 1680 bp in size. The reaction mixture contains 5 pmol of forward and reverse primers, 100 μM dNTPs, 1x Taq DNA polymerase buffer and 2 U Taq DNA polymerase (Promega) in a total volume of 50 μl. The initial denaturation step is 94°C for 2 min, followed by 29 cycles of 94°C for 30 sec, 60°C for 60 sec and 72°C for 90 sec. Cycling is completed by a final elongation step at 72°C for 5 min. The amplification products are analyzed by electrophoresis in a 1% agarose gel and visualized with a fluorescent stain. The gluA_short fragment is amplified using primers EsAg5f: 5′-TATCAGATCTACCCGCGC-3′ and EsAg5_5r: 5′-TTGATGCCAAGCTGTTGC-3′ resulting in a 105-bp amplicon. The PCR cycling conditions are as described above except annealing is at 62°C for 30 sec and extension at 72°C for 30 sec. The amplification products are analyzed in a 2% agarose gel. 26.2.2.2 Real-Time PCR Identification A real-time PCR assay targeting the dnaG gene, which is a component of the macromolecular synthesis operon, was originally developed by Seo and Brackett.59 The method was modified by Drudy et al.48 using the TaqMan probe (6-FAMACAGAGTAGTAGTTGTAGAGGCCGTGCTTCC-TMR). To confirm identification of E. sakazakii, total genomic DNA is purified from presumptive colonies as outlined

above. The following primers are used for amplification: Fwd: 5′-GGGATATTGTCCCCTGAAACAG-3′ and Rev: 5′-CAGGAATAAGCCGCGATT-3′.59 Amplification of the target region is performed in a final volume of 20 μl, using 200 μM deoxynucleoside triphosphates, 4.0 mM MgCl2, 2.0 μl of 10 × reaction buffer, 10 mM of each primer, 2.5 mM of probe, 1 U Taq DNA polymerase, and 2 µl DNA template (containing approximately 100 ng of DNA). The recommended reaction conditions are: 95°C for 15 min followed by 40 cycles of 95°C for 15 sec and 60°C for 50 sec. In the study by Drudy et al.48 the RT-PCR was performed on a Corbett Research Rotor-Gene RG 3000 instrument.

26.3 Conclusions and Future Perspectives To date, there are only two molecular identification methods for E. sakazakii for which extensive evaluation data is publically available. Both methods are designed to target all species of the newly described genus Cronobacter (E. sakazakii) and both methods perform equally well in terms of sensitivity and specificity. Considering the new taxonomic description for Cronobacter, there is perhaps a need for the development of methods for the species-specific identification of organisms within this genus. This can be facilitated by the publication of the complete E. (Cronobacter) sakazakii genome sequence, as well as the genome sequence of a second type strain—Cronobacter turicensis 3032—which will be available in the near future. Genomic analysis will provide deeper insights into the genetic organization of Cronobacter species and will enable the identification of potential targets for molecular methods that will allow accurate discrimination of species. An interesting approach into the direction of molecular based serotyping of E. sakazakii was published recently by Mullane et al.78 In this study nucleotide polymorphism in the O-antigen coding locus rfb was determined and a PCR assay targeting the genes specific for the two prominent serotypes in E. sakazakii was developed. Last but not least, there are several efforts undergoing to improve the cultural detection method for Cronobacter spp., which might also lead to an improvement in the performance and application of the molecular identification methods.

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5. Farmer, J.J. et al. Enterobacter sakazakii: a new species of Enterobacteriaceae isolated from clinical species. Int. J. Syst. Evol. Bacteriol., 30, 569, 1980. 6. Lehner, A., Tasara, T. and Stephan, R. 16S rRNA gene based analysis of Enterobacter sakazakii strains from different sources and development of a PCR assay for identification. BMC Microbiol., 4, 43, 2004. 7. Iversen, C., Waddington, M. and Forsythe, S.J. Identification and phylogeny of Enterobacter sakazakii relative to Enterobacter and Citrobacter species. J. Clin. Microbiol., 42, 5368, 2004. 8. Iversen, C. et al. The taxonomy of Enterobacter sakazakii: proposal of a new genus Cronobacter gen. nov., and descriptions of Cronobacter sakazakii comb. nov., Cronobacter sakazakii subsp. sakazakii, comb. nov. Cronobacter sakazakii subsp. malonaticus subsp. nov., Cronobacter turicensis sp. nov., Cronobacter muytjensii sp. nov. Cronobacter dublinensis sp. nov. and Cronobacter genomospecies 1. BMC Evol. Biol., 7, 64, 2007. 9. Iversen, C. et al. Cronobacter gen. nov., a new genus to accommodate the biogroups of Enterobacter sakazakii, and proposal of Cronobacter sakazakii gen. nov. comb. nov., C. malonaticus sp. nov., C. turicensis sp. nov., C. muytjensii sp. 3 nov., C. dublinensis sp. nov., Cronobacter genomospecies 1, and of three subspecies, C. dublinensis sp. nov. subsp. dublinensis subsp. nov., C. dublinensis sp. nov. subsp. lausannensis subsp. nov., and C. dublinensis sp. nov. subsp. lactaridi subsp. nov. Int. J. Syst. Evol. Microbiol., 58, 1442, 2008. 10. Iversen, C. et al. Identification of Cronobacter spp. (Enterobacter sakazakii). J. Clin. Microbiol., 45, 3814, 2007b. 11. Anonymous, FAO-WHO. Enterobacter sakazakii and other microorganisms in powdered infant formula: meeting report. MRA Series 6, WHO, Geneva, 2004. 12. Anonymous, FAO-WHO. Enterobacter sakazakii and Salmonella in powdered infant formula, Second Risk Assessment Workshop. 16–-20th January, WHO Rome, Italy, 2006. 13. Anonymous, Technical Specification ISO/TS 22964. 2006. Milk and milk products – detection of Enterobacter sakazakii. ISO/TS 22964:2006(E) and IDF/RM 210:2006(E).210, 2006(E), First Edition. 14. Simmons, B.P. et al. Enterobacter sakazakii infections in neonates associated with intrinsic contamination of powdered infant formula. Infect. Control Hosp. Epidemiol., 10, 398, 1989. 15. Clark, N.C. et al. Epidemiologic typing of Enterobacter sakazakii in two neonatal nosocomial outbreaks. Diagn. Microbiol. Infect. Dis., 13, 467, 1990. 16. Van Acker, J. et al. Outbreak of necrotizing enterocolitis associated with Enterobacter sakazakii in powdered milk formula. J. Clin. Microbiol., 39, 293, 2001. 17. Mullane, N. et al. Enterobacter sakazakii: and emerging bacterial pathogen with implications for infant health. Minerva Pediatr., 59, 137, 2007. 18. Muytjens, H.I. et al. Analysis of eight cases of neonatal meningitis and sepsis due to Enterobacter sakazakii. Appl. Environ. Microbiol., 70, 5692, 1983. 19. Nazarowec-White, M. and Farber, J.M. Enterobacter sakazakii: a review. Int. J. Food Microbiol., 34, 103, 1997. 20. Lehner, A. and Stephan, R. Microbiological, epidemiological, and food safety aspects of Enterobacter sakazakii. J. Food Prot., 67, 2850, 2004. 21. Adamson, D.H. and Rogers, J.R. Enterobacter sakazakii meningitis with sepsis. Clin. Microbiol. Newslett., 3, 19, 1981.

Molecular Detection of Foodborne Pathogens 22. Kleimann, M.B. et al. Meningoencephalitis and compartmentalization of the cerebral ventricles caused by Enterobacter sakazakii. J. Clin. Microbiol., 14, 352, 1981. 23. Bowen, A.B. and Braden, C.R. Invasive Enterobacter sakazakii disease in infants. Emerg. Infect. Dis., 12, 1185, 2006. 24. Willis, J. and Robinson, J.E. Enterobacter sakazakii meningitis in neonates. Pediatr. Infect. Dis. J., 7, 196, 1988. 25. Urmenyi, A.M.C. and Franklin, A.W. Neonatal death from pigmented coliform infection. Lancet, 1, 313, 1961. 26. Muytjens, H.L. and Kollee, L.A.A. Enterobacter sakazakii meningitis in neonates: causative role of formula. Pediatr. Infect. Dis. J., 9, 372, 1990. 27. Bar-Oz, B. et al. Enterobacter sakazakii infection in the newborn. Acta Paediatr., 90, 356, 2001. 28. J. Levy, J. et al. Contaminated enteral nutrition solutions as a cause of nosocomial bloodstream infection: a study using plasmid fingerprinting. J. Parent. Enter. Nutr. 13, 228, 1989. 29. Oie, S. and Kamiya, A. Comparison of microbial contamination of enteral feeding solution between repeated use of administration sets after washing with water and after washing followed by disinfection. J. Hosp. Infect., 48, 304, 2001. 30. Noriega, F.R. et al. Nosocomial bacteremia caused by Enterobacter sakazakii and Leuconostoc mesenteroides resulting from extrinsic contamination of infant formula. Pediatr. Infect. Dis., 9, 447, 1990. 31. Smeets, L.C. et al. Genetische karakterisatie van Enterobacter sakazakii-isolaten van Nederlandse patiënten met neonatale meningitis. Ned. Tijdschr. Med. Microbiol.. 6, 113, 1998. 32. Biering, G. et al. Three cases of neonatal meningitis caused by Enterobacter sakazakii in powdered milk. J. Clin. Microbiol., 27, 2054, 1989. 33. Nazarowec-White, M. and Farber, J.M. Phenotypic and genotypic typing of food and clinical isolates of Enterobacter sakazakii. J. Med. Microbiol., 48, 559, 1999. 34. Townsend, S., Hurrell, E. and Forsythe, S. Virulence studies of Enterobacter sakazakii isolates associated with a neonatal intensive care unit outbeak. BMC Microbiol., 8, 64, 2008. 35. Lucas, A., and Cole, T.J. Breast milk and neonatal necrotizing enterocolitis. Lancet, 336, 1519, 1990. 36. Chan, K.L. et al. A study of preantibiotic bacteriology in 125 patients with necrotizing enterocolitis. Acta Paediatr. (Suppl.), 396, 45, 1994. 37. Anonymous, U.S. Food and Drug Administration. Isolation and enumeration of Enterobacter sakazakii from dehydrated powdered infant formula, 2002. [Online] http://www.cfsan. fda.gov~comm/mmesakaz.html. 38. Guillaume-Gentil, O. et al. A simple and rapid cultural method for detection of Enterobacter sakazakii in environmental samples. J. Food Prot., 68, 64, 2005. 39. Iversen, C., Druggan, P. and Forsythe. S.J. A selective differential medium for Enterobacter sakazakii. Int. J. Food Microbiol., 96, 133, 2004. 40. Leuschner, R.G.K. and Bew, J. A medium for the presumptive detection of Enterobacter sakazakii in infant formula: interlaboratory study. J. AOAC Int., 87, 604, 2004. 41. Oh, S.W. and Kang, D.H. Fluorogenic selective and differential medium for isolation of Enterobacter sakazakii. Appl. Environ. Microbiol., 70, 5692, 2004. 42. Restaino, L. et al. A chromogenic plating medium for the isolation and identification of Enterobacter sakazakii from foods, food ingredients and environmental sources. J. Food Prot., 69, 315, 2006.

Enterobacter 43. Song, K.Y. et al. Evaluation of a chromogenic medium supplemented with glucose for detecting Enterobacter sakazakii. Microbiol. Biotechnol., 18, 579, 2008. 44. Lehner, A. et al. Comparison of two chromogenic media and evaluation of two molecular based identification systems for Enterobacter sakazakii detection. BMC Microbiol., 6, 15, 2006. 45. Iversen, C. and Forsythe, S. Comparison of media for the isolation of Enterobacter sakazakii. Appl. Environ. Microbiol., 73, 48, 2007. 46. Iversen, C. et al. Development of a novel screening method for the isolation of Cronobacter spp. (Enterobacter sakazakii). Appl. Environ. Microbiol., 74, 2550, 2008. 47. Mullane, N.R. et al. Detection of Enterobacter sakazakii in dried infant milk formula by cationic magnetic bead capture. Appl. Environ. Microbiol., 72, 6325, 2006. 48. Drudy, D. et al. Characterization of a collection of Enterobacter sakazakii isolates from environmental and food sources. Int. J. Food Microbiol., 110, 127, 2006. 49. Witthuhn, R.C., Kemp, F. and Britz, T.J. Isolation and PCR detection of Enterobacter sakazakii in South African food products, specifically infant formula milks. World J. Microbiol. Biotechnol., 23, 151, 2007. 50. Hassan, A.A. et al. Characterization of the gene encoding the 16S rRNA of Enterobacter sakazakii and development of a species-specific PCR method. Int J. Food Microbiol., 116, 214, 2007. 51. Nair Mohan. K.M. and Ventkitanarayanan K.S. Cloning and sequencing of the ompA gene of Enterobacter sakazakii and development of an ompA targeted PCR for rapid detection of Enterobacter sakazakii in infant formula. Appl. Environ. Microbiol., 72, 2539, 2006. 52. Lehner, A. et al. Molecular characterization of the alpha glucosidase activity in Enterobacter sakazakii reveals the presence of a putative gene cluster for palatinose metabolism. Syst. Appl. Microbiol., 29, 609, 2006. 53. Kothary, M.M. et al. Characterization of the zinc-containing metalloprotease encoded by zpx and development of a species-specific detection method for Enterobacter sakazakii. Appl. Environ. Microbiol., 73, 4142, 2007. 54. Malorny, B. and Wagner, M. Detection of Enterobacter sakazakii strains by real-time PCR. J. Food Prot., 68, 1623, 2005. 55. Lehmacher, A., Fiegen, M. and Hansen, B. Real-time PCR von Enterobacter sakazakii in Säuglingsanfangsnahrung. J. Verbr. Lebensm., 2, 218, 2007. 56. Kang S.E., Nam, Y.S. and Hong, K.W. Rapid detection of Enterobacter sakazakii using TaqMan real-time PCR assay. Microbiol. Biotechnol., 17, 526, 2007. 57. Liu, Y. et al. Real time PCR using TaqMan and SYBR Green for detection of Enterobacter sakazakii in infant formula. J. Microbiol. Methods, 65, 21, 2006. 58. Derzelle, S. and Dilasser, F. A robotic DNA purification protocol and real-time PCR for the detection of Enterobacter sakazakii in powdered milk formulae. BMC Microbiol., 6, 100, 2006. 59. Seo, K.H. and Brackett, R.E. Rapid, specific detection of Enterobacter sakazakii in infant formula using a real time PCR assay. J. Food Prot., 68, 59, 2005. 60. Mullane, N.R. et al. Application of pulsed-field gel ­electrophoresis to characterise and trace the prevalence of Enterobacter sakazakii in an infant formula processing facility. Int. J. Food Microbiol., 116, 73, 2007.

367 61. Bruce, J. Automated system rapidly identifies and characterizes microorganisms in food. Food Technol., 50, 77, 1996. 62. Clementino, M.M. et al. PCR analyses of tRNA intergenic spacer, 16S–23S internal transcribed spacer, and randomly amplified polymorphic DNA reveal inter and intraspecific relationships of Enterobacter cloacae strains. J. Clin. Microbiol., 39, 3865, 2001. 63. Babalola, O.O. et al. Characterization of potential ethyleneproducing rhizosphere bacteria of Striga-infested maize and sorghum. Afr. J. Biotechnol., 1, 67, 2003. 64. Sanjaq, S. et al. Improvement and validation of RAPD in comparison to PFGE analysis of Enterobacter sakazakii strains. Int. J. Med. Microbiol., 297, 97, 2007. 65. Kim, K. et al. Prevalence and genetic diversity of Enterobacter sakazakii in ingredients of infant foods. Int. J. Food Microbiol., 122, 196, 2008. 66. Proudy, I. et al. Genotypic characterization of Enterobacter sakazakii isolates by PFGE, BOX-PCR and sequencing of the fliC gene. Appl. Environ. Microbiol., 104, 26, 2008. 67. Proudy, I. et al. Tracing of Enterobacter sakazakii isolates in infant milk formula processing by BOX-PCR genotyping. J. Appl. Microbiol., 2008. 68. Healy, B. et al. Evaluation of an automated rep-PCR system for subtyping Enterobacter sakazakii. J. Food Prot., 71, 1372, 2008. 69. Keim, P. et al. Multiple-locus variable–number tandem-repeat analysis reveals genetic relationship within Bacillus anthracis. J. Bacteriol., 182, 2928, 2000. 70. Le Fleche, P. et al. A tandem repeats database for bacterial genomes: application to the genotyping of Yersinia pestis and Bacillus anthracis. BMC Microbiol., 1, 2, 2001. 71. Lindstedt, B.A. et al. DNA fingerprinting of Shiga-toxin producing Escherichia coli O157 based on multiple-locus variable-number tandem-repeats analysis (MLVA). Ann. Clin. Microbiol. Antimicrob., 2, 12, 2003. 72. Lindstedt, B.A. et al. Multiple-locus variable-number tandem-repeats analysis of Salmonella enterica subsp. enterica serovar Typhimurium using PCR multiplexing and multicolor capillary electrophoresis. J. Microbiol. Methods, 59, 163, 2004. 73. Mullane, N.R. et al. Development of multiple-locus variable tandem repeat analysis for the molecular subtyping of Enterobacter sakazakii. Appl. Environ. Microbiol., 74, 1223, 2008. 74. Vos, P. et al. AFLP: a new technique for DNA fingerprinting. Nucleic Acids Res., 23, 4407, 1995. 75. Mueller, U.G. and Wolfenbarger, L.L. AFLP genotyping and fingerprinting. Tree, 14, 389, 1999. 76. Rademaker, J.L. et al. Comparison of AFLP and rep-PCR genomic fingerprinting with DNA-DNA homology studies: Xanthomonas as a model system. Int J. Syst. Evol. Microbiol., 50, 665, 2000. 77. Mougel, B. et al. A mathematical method for determining genome divergence and species delineation using AFLP. Int. J. Syst. Evol. Microbiol, 52, 573, 2002. 78. Mullane, N. et al. Molecular analysis of Enterobacter ­sakazakii O-antigen gene locus. Appl. Environ. Microbiol., 74, 3783, 2008. 79. Institut de Veille Sanitaire. Infections à Enterobacter sakazakii associées à la consommation d’une préparation en poudre pour nourrissons. France, octobre à décembre 2004. Rapport d’investigation, 2006.

27 Escherichia

Devendra H. Shah, Smriti Shringi, Thomas E. Besser, and Douglas R. Call Washington State University

Contents 27.1 Introduction.................................................................................................................................................................... 369 27.1.1 Occurrence and Epidemiology......................................................................................................................... 369 27.1.2 Pathogenesis and Virulence Traits.................................................................................................................... 370 27.1.3 Conventional Diagnostic Techniques................................................................................................................ 371 27.1.3.1 Culture and Isolation........................................................................................................................ 371 27.1.3.2 Latex Agglutination Test.................................................................................................................. 373 27.1.3.3 Enrichment and Immunomagnetic Separation................................................................................ 373 27.1.3.4 ELISA.............................................................................................................................................. 374 27.1.4 Molecular Diagnostic Techniques.................................................................................................................... 375 27.1.4.1 PCR.................................................................................................................................................. 375 27.1.4.2 Real-Time PCR................................................................................................................................ 377 27.1.4.3 Reverse Transcriptase PCR............................................................................................................. 380 27.1.4.4 Microarray....................................................................................................................................... 380 27.2 Methods.......................................................................................................................................................................... 381 27.2.1 Reagents and Equipment................................................................................................................................... 382 27.2.2 Sample Preparation .......................................................................................................................................... 382 27.2.3 Detection Procedure......................................................................................................................................... 382 27.3 Concluding Remarks...................................................................................................................................................... 383 Acknowledgments...................................................................................................................................................................... 383 References.................................................................................................................................................................................. 383

27.1  Introduction Escherichia coli O157:H7 (hereafter referred to as O157:H7) is an important foodborne pathogen and a causative agent of diarrhea, hemorrhagic colitis (HC) and a life threatening post-diarrheal sequela called hemolytic uremic syndrome (HUS).1,2 The genus Escherichia covers a number of Gramnegative, non-spore forming bacterial species in the family Enterobacteriaceae, of which Escherichia coli is one of the most numerous aerobic commensal species in the gastrointestinal tracts of humans and animals. While many E. coli strains are harmless, some, such as serotype O157:H7, can cause serious illnesses in humans. Clinically, HC is characterized by severe abdominal pain and bloody diarrhea. Approximately 10–15% of children below 10 years of age with HC progress to HUS, which is characterized by hemolytic anemia, thrombocytopenia, oliguria-anuria, acute renal failure and, rarely, seizures.3,4 In general, immunocompromised individuals, the very young and very elderly individuals are more susceptible to severe disease or death following O157:H7 infection. Ingestion of contaminated food and water is the major source of infection, but direct contact with infected humans or animals is also known to transmit infection to susceptible individuals.5 Serotype O157:H7 belongs to a group of diarrheagenic E. coli commonly referred as enterohemorrhagic E. coli (EHEC).

EHEC strains cause disease due to their ability to attach intimately to the intestinal mucosa and cause attaching-effacing lesions and also by their ability to produce Shiga-like toxins (Stx) that are responsible for inducing HC and HUS. Therefore, EHEC strains are also classified as subset of Stx producing E. coli (STEC) that are pathogenic to humans. While STEC is the term used for all Shiga toxin ­producing E. coli, it is not considered a defined E. coli pathovar.2 Nevertheless, the terms EHEC and STEC have been used interchangeably in the literature.

27.1.1 Occurrence and Epidemiology More than 100 serotypes of Shiga-toxin producing E. coli are associated with human infections6 and resultant illness is similar to that of O157:H7 infection.1,7 Nonetheless, O157:H7 is the most frequently isolated serotype from cases of bloody diarrhea and HUS.5 E. coli O157:H7 was first isolated in the US in 1982 during an outbreak of HC in which 47 individuals ingested contaminated hamburgers at a fast food chain in Michigan and Oregon.8 A retrospective examination of more than 3,000 isolates obtained between 1973 and 1982 revealed only one isolate of O157:H7 that was originally recovered from a 50-year-old woman who had experienced acute bloody diarrhea in 1975.8 Thereafter, sporadic cases 369

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as well as outbreaks due to O157:H7 infection have been reported increasingly in the US, UK, Europe, and other developed parts of the world.9–15 Between 1982 and 2002, a total of 350 outbreaks of O157:H7 infections were reported from 49 states in the US, representing 8,598 cases; resulting in 1,493 (17%) hospitalizations, 354 (4%) HUS cases, and a total of 40 (0.5%) deaths.5 In 2006, the Center for Disease Control FoodNet surveillance program reported 590 cases of O157:H7 infection in humans with an overall annual incidence of 1.31 cases per 100,000 people in the US.16 In the UK, the Health Protection Agency has reported that there were 70,603 food poisoning notifications in 2006. Of these, 1,003 (1.4%) were caused by O157:H7.15 The majority of outbreaks in North America are caused by typical O157:H7 isolates that can be distinguished from most of the other E. coli by their inability to rapidly (within 24 h) ferment sorbitol and by their inability to produce β-glucuronidase. There are also atypical strains that are non-motile (O157:NM) or that can rapidly ferment sorbitol and these have been associated with HUS and HC in Germany and other European countries.17,18 There has been a recent increase in the reported incidence of human infections associated with non-O157 STEC. A survey in the US found that a total of 940 non-O157 STEC were isolated from individuals with sporadic illnesses between 1982 and 2002.7 Out of 76 different serogroups isolated from these outbreaks, the most common O-antigen types included O26 (22%), O111 (16%), O103 (12%), O121 (8%), O45 (7%) and O145 (5%). In Europe, non-O157 serotypes are more commonly isolated than O157:H7.10,19–21 Recent reports from certain parts of the US (Montana, Washington, and Connecticut) show that non-O157 STEC are isolated at a frequency similar to that of O157:H7.22–24 In general, however, the relative isolation rates of non-O157 isolates vary from study to study and are influenced by geographical areas and methods employed for sampling and testing. Unlike O157:H7, the incidence, trends, and epidemiology of pathogenic non-O157 STEC are not well understood.7 It is also important to note that most strains of non-O157 STEC do not cause human illness, and some nonO157 STEC isolates recovered from diarrheal stool may not be pathogens.7,25 Moreover, pathogenic non-O157 E. coli are often biochemically indistinguishable from nonpathogenic E. coli and the identification of these bacteria requires complex biochemical, serological, and molecular testing that are not routinely performed by clinical microbiology laboratories. Contaminated food is one of the most important sources of O157:H7 infection, although contaminated water or direct contact with infected humans or animals are also reported as vehicles for transmission of infection.5 O157:H7 is ubiquitous on cattle operations in North America and has been isolated from the majority, if not all, operations studied longitudinally26,27 and estimates of point-in-time prevalence at the herd level have been as high as 63–100%.28–30 Fecal testing of cattle worldwide revealed a wide range of prevalence rates for O157:H7 in both dairy (0.2–48.8%) and beef (0.2–27.8%) cattle operations.31,32 Global assessments of contamination rates of O157:H7 in retail beef products during the last three decades showed that the prevalence of O157:H7 ranged from

Molecular Detection of Foodborne Pathogens

0.1 to 54.2% in ground beef, 0.1–44% in sausage, 1.1–36% in various retail cuts, and 0.01–43.4% in whole carcasses,25 suggesting that the high contamination rate of beef products corresponds to the wide-spread prevalence in cattle operations. A summary of O157:H7 associated outbreak investigations between 1982 and 2002 revealed that contaminated food accounted for 183 (52%) out of 350 outbreaks and 61% of 8,598 outbreak related cases, respectively, and that the contaminated beef products were reported as the primary source of infection in the majority (86 out of 183) of food related outbreaks.5 While there is a link between prevalence in cattle operations, contamination of retail beef products and subsequent infection of humans, the ecology and epidemiology of O157:H7 in cattle are complex and remain poorly understood.33–37 Although the majority of O157:H7 illnesses are related to consumption of contaminated beef, several outbreaks have been associated with other food commodities including lettuce, apple cider, salad, coleslaw, melons, sprouts, grapes, cheese, butter, and raw milk.4,5 Out of 350 outbreaks reported in the US between 1982 and 2002, direct contact with infected animals or humans and contaminated water (both recreational and drinking) were identified as a source of infection in 93 (26.5%) outbreaks, whereas the source of infection could not be traced in 116 (33%) outbreaks,5 indicating a complex ecology, and distribution O157:H7 in the environment and foods. Due to the ubiquitous nature of O157:H7, the development of portable and robust methods for its detection from different food and environmental matrices remains a challenging task. Consequently, most of the efforts in last three decades have been primarily directed towards the development of rapid and sensitive methods (both conventional and molecular) for the detection of O157:H7 from different types of foods and environment.

27.1.2  Pathogenesis and Virulence Traits O157:H7 is highly infectious for human beings with an estimated infectious dose being very low (~10–100 organisms).38 Unfortunately, there are no animal models that completely reflect O157:H7 infections in humans and thus, mechanisms of pathogenesis are not completely understood. Nevertheless, several virulence traits have been identified and their roles in the causation of disease have been elucidated. One important feature of O157:H7 infection is its ability to attach intimately to the intestinal epithelium and cause intestinal attaching and effacing (A/E) lesions. The other hallmark of O157:H7 pathogenicity is its ability to cause systemic effects via one or more Shiga-like toxins (Stx1, Stx2, and their variants). The genes encoding Stx1 and Stx2 are located on lambdoid bacteriophages that are integrated in the genome of O157:H7.39,40 Surveys have shown that the vast majority of North American O157:H7 isolates associated with HUS possess Stx2 alone or in combination with Stx1 and that only a small fraction possess Stx1 alone.41,42 Stx are composed of 5 B subunits and a single A subunit. The B subunits bind to glycophospholipid receptors (Gb3) on the surface of eukaryotic cells and the A subunit is an N-glycosidase, which inhibits protein synthesis and disrupts the large eukaryotic ribosomal subunit.43 These

Escherichia

toxins bind to and damage endothelial cells in the intestine, kidney, and brain.2,3 Although Stx1 is relatively homogeneous, two subtypes (Stx1c and Stx1d) have been described.44,45 Five subtypes of Stx2 have been identified, and these include Stx2c, Stx2d, Stx2e, Stx2f, and Stx2g.43,46 Epidemiological studies have indicated that stx2 is more associated with severe human disease than stx147 and among stx2 variants, stx2, and stx2c are frequently found in strains isolated from patients with HUS, while strains producing stx2d are usually isolated from cases of uncomplicated diarrhea.48 Typical O157:H7 strains possess other virulence factors encoded by the chromosomally located enterocyte effacement (LEE) pathogenicity island (PAI) and a large (~90 kb) virulence plasmid referred to as pO157. The LEE is integrated adjacent to either selC, pheV or pheU tRNA loci and consists of three functional modules.39,40 The first set (sepA to sepI) encode a type III secretion system (TTSS) that exports effector molecules. The second set (espA to espD) encode structural proteins of the TTSS, and the third set (eaeA and tir) encode the outer membrane adhesin, intimin, and its receptor Tir, which is translocated into the host cell plasma membrane via the TTSS apparatus.49 The LEE enables the bacteria to efficiently colonize the intestine of the host by binding between intimin, the product of eae gene, and its receptor Tir. Binding of LEE positive bacteria to the intestinal cells results in a characteristic A/E phenotype.3,50 The C′ terminus of the eaeA gene is highly variable. Based on the sequence and antigenic differences in the C′ terminus of eaeA several distinct types of intimin (α, β, γ, δ, ζ, η, θ, ι, and κ) have been identified of which γ intimin is generally produced in O157:H7 and several other non-O157 serotypes.51 In vitro studies that test adherence to cultured cells have identified a non-LEE adherence related gene, iha.52 In addition, the pO157 plasmid harbors several genes that are suspected to play a role in disease. These genes include hlyA (encoding hemolysin), katP (encoding catalase-peroxidase), espP (encoding serine protease) and toxB.3,53 The hlyA and toxB gene are present in almost all isolates of O157:H7 while the katP and espP genes can be detected in two thirds of the isolates.53 Additionally, the genome of O157:H7 contains 16 loci encoding genes involved in biosynthesis of other putative adhesins including fimbriae or pili.39,40 Currently, it is unclear how many of these are functional and necessary to induce disease. Several other (PAIs) and putative virulence associated genes have been described,54 but their role in pathogenesis of O157:H7 is not fully understood. Regardless of their functional status, these real or putative virulence-related genes provide targets for development of rapid, sensitive, and specific molecular methods for O157:H7, including assays based on PCR and various hybridization platforms.

27.1.3 Conventional Diagnostic Techniques 27.1.3.1  Culture and Isolation Conventional methods for isolation of O157:H7 include a combination of selective plating, enrichment procedures, and immunomagnetic separation (IMS) that is usually followed by confirmation via biochemical and serological

371

testing (Figure 27.1). Typical O157:H7 strains differ from most other E. coli strains in several ways. Over 95% of all E. coli strains rapidly (within 24 h) ferment sorbitol and produce β-glucuronidase, whereas typical O157:H7 strains do not ferment sorbitol within 24 h and fail to produce functional β-glucuronidase.55–58 O157:H7 strains do not ferment rhamnose on agar plates whereas 60% of nonsorbitol-fermenting E. coli strains belonging to serogroups other than O157:H7 will ferment rhamnose.59 These features have been exploited to develop sorbitol-MacConkey’s agar (SMAC) for isolation and differentiation of O157:H7 strains from several other serogroups of E. coli.56,60 The selectivity of SMAC was later improved by the addition of sub-inhibitory concentrations of cefixime (0.05 µg/ml) and 0.5% (w/v) rhamnose to make CR-SMAC agar,61 and cefixime and potassium tellurite to make CT-SMAC, agar.62 SMAC, CR-SMAC, and CT-SMAC media contain 1% sorbitol instead of 1% lactose with the standard MacConkey’s medium base. Cefixime was added to inhibit the growth of Proteus spp. and tellurite was added to inhibit the growth of Providentia, Aeromonas, Morganella, and Plesiomonas spp., which are prevalent in the feces of cattle and humans. The majority of E. coli of the fecal flora can ferment sorbitol within 24 h and yield pink colonies, whilst nonsorbitol fermenting E. coli O157 produce pale or colorless colonies on CT-SMAC agar. CT-SMAC is by far the most commonly used selective plating media for isolation of O157:H7. This system is not perfect, however, as false positives are possible from non-O157 isolates, and tellurite-sensitive and rapidly sorbitol-fermenting atypical strains of O157:H7 have been reported and thus will not be detected by these media.63 Moreover, colony characteristics of many organisms other than O157:H7, especially other sorbitol nonfermenting serogroups of E. coli, Proteus spp., Morganella spp., and Hafnia spp., appear identical to O157:H7 when grown on CT-SMAC. Several other chromogenic/fluorogenic selective media have also been developed including Rainbow agar O157 (RB O157; Biolog Inc., Hayward, CA), Biosynth culture media O157:H7 (BCM O157:H7; Biosynth Staad, Switzerland), Fluorocult O157:H7 (HC, Merck, Darmstadt, Germany), CHROMagar O157 (CHROMagar, Paris, France), and O157:H7 ID-F agar (O157 H7 ID-F; bioMerieux SA, Marcyl’Etoile, France). The efficacy of these media has been tested for the isolation and differential detection of O157:H7 from foods as well as clinical samples. RB O157 capitalizes on the inability of O157:H7 to produce β-glucuronidase. The addition of selective agents for E. coli and chromogenic substrates for β-glucuronidase and β-galactosidase allows selective detection. Most O157:H7 isolates are glucuronidase-negative and galactosidase-positive and their colonies appear as black or gray on this medium, whereas commensal E. coli strains produce β-glucuronidase and hence appear as pink colonies. Some non-O157 strains overproduce β-galactosidase relative to β-glucuronidase and produce intermediate (purple or magenta) colored colonies. One study examined 585 isolates of E. coli including typical and atypical strains of O157:H7 and several non-O157 STEC serogroups isolated from clinical

372

Molecular Detection of Foodborne Pathogens

Feces/food/environmental samples

Selective enrichment (6–24 h) Immunomagnetic separation (1–2 h)

ELISA (Stx and/or O157, H7) Direct plating on CT-SMAC and/or any other chromogenic medium (16–24 h) No O157:H7-like colonies Possible false negative Go to enrichment step

Prepare DNA for PCR (15–30 min)

Typical O157:H7 colonies

Serotyping and biochemical typing

PCR or/and microarray (2–4 h) (rfbE, fliC, stx1, stx2, eaeA, uidA, and hlyA)

rfbE+, fliC+, stx1+, stx2+, eaeA+, uidA+, and hlyA+ rfbE+, fliC+, stx1–, stx2+, eaeA+, uidA+, and hlyA+ rfbE+, fliC+, stx1+, stx2–, eaeA+, uidA+, and hlyA+ Typical or atypical O157:H7

rfbE–, fliC–, stx1±, stx2±, eaeA±, uidA–, and hlyA± Non-O157 E. coli?

rfbE+, fliC+, stx1–, stx2–, eaeA+, uidA+, and hlyA+ nonpathogenic O157:H7?

rfbE–, fliC+, stx1±, stx2±, eaeA±, uidA–, and hlyA± Other E. coli?

Proceed for molecular fingerprinting (e.g., PFGE)

Proceed for further confirmation by biochemical and serological testing

rfbE+, fliC–, stx1±, stx2±, eaeA±, uidA–, and hlyA± Other O157 E. coli?

Figure 27.1  A schematic of tiered microbiological and PCR scheme used to detect virulence factors and serotype-specific markers of O157:H7 from different sample matrices.

and environmental sources from Africa, Asia, Australia, Europe, and North America.64 This work showed that, in addition to O157:H7 isolates, several non-O157 STEC strains also produced a characteristic black colonies on RB agar and were indistinguishable from O157:H7 strains.64 Some nonO157 STEC strains produced characteristic mauve, red or pink colonies and were distinguishable from non-shiga toxin producing E. coli. In a separate study, Bettelheim65 used a collection of 585 isolates of E. coli to test the reliability of CHROMagar for the differential detection of O157:H7 and observed that although >90% of O157:H7/NM strains could be readily isolated and recognized by the characteristic pink colored colonies, several non-O157 STEC also produced similar colonies and were indistinguishable from O157:H7/ NM isolates. Reliability of a recently available ID-F agar, which contains carbohydrates, two chromogenic substrates to detect β-galactosidase and β-glucuronidase, and sodium deoxycholate to increase selectivity of Gram-negative rods, was also tested for the differential detection of O157:H7/NM strains.66 Out of 63 O157:H7/NM strains, 59 (93.7%) strains produced characteristic green colored colonies and out of the four O157:NM strains that failed to produce typical colonies, three were identified as Shiga-toxin negative. In addition, three non-O157 strains including one O55:H7 also produced

green colored colonies not distinguishable from O157:H7, whilst 284 non-O157 STEC or Shiga-toxin negative E. coli produced purple colonies and could be readily distinguished from O157:H7/NM strains.66 Recently, the performance of CHROMagar was compared with that of SMAC agar for the specific detection of O157:H7 from clinical specimens.67 A total of 3,116 stool specimens were tested, of which 27 generated culture positive results for O157:H7. CHROMagar was reported to have a higher sensitivity (96.3%) and negative predictive value (100%) than SMAC while specificity (100%) and positive predictive values (100%) were similar for both media.67 These authors also reported that the diagnostic efficiency of CHROMagar was better than SMAC due to the significant reduction in the numbers of false positive colony picks needed for further confirmation by serological and biochemical testing, thereby reducing labor and material costs.67 In another study, reliability of four chromogenic/fluorogenic media (RB, BCM, Fluorocult HC and SMAC) was evaluated for differential detection of O157:H7 from pure cultures and food samples such as ground beef and raw milk.68 Examination of 34 reference strains of O157:H7/NM showed false negative results with BCM (3%), RB (8.8%), HC, and SMAC (5.9%), whereas one out of 12 (8.3%) non-O157 E. coli strains generated false

Escherichia

positive results with BCM, RB, and SMAC, but no false positives were detected with HC agar.68 Examination of 60 food samples led to isolation of 466 Gram-negative bacteria of different genera including non-O157 E. coli, of which 57.3%, 6.2%, 3.3%, and 2.1% isolates produced false positive results on SMAC, HC, BCM, and RB agar, respectively.68 Collectively, studies of efficiency of different media for the differential detection of O157:H7 have indicated that although currently available plating media provide a valuable tool for the specific and sensitive detection of O157:H7, but the false positive as well as false negative results may be produced and therefore, further testing of the suspect colonies using conventional serological and biochemical typing is generally required for definitive identification. 27.1.3.2 Latex Agglutination Test (LAI) Confirmation of E. coli O157:H7 requires testing for the O157 somatic and H7 flagellar antigens. Several latex agglutination test (LAT) kits are commercially available. These are composed of latex particles coated with antiserum against O157 and H7 antigens for screening and detection of suspect colonies of O157:H7. Evaluation of performance of three commercial LATs for the specific detection of O157 and one LAT for the specific detection of H7 antigens revealed 100% correlation of commercial O157 kits with CDC reference antisera.69 The H7 reagent had a diagnostic sensitivity of 96% and diagnostic specificity of 100% compared with the CDC antisera.69 A study that compared four commercial LAT kits for their efficacy to detect O157 and H7 antigens revealed diagnostic sensitivities and specificities of 99–100%, respectively, for all kits.70 Nevertheless, special precautionary measures have been suggested with the use of these LAT test kits. False positive interpretations could arise if the latex controls were not routinely used or due to the cross reactivity of O157 antisera with strains of Citrobacter freundii and Salmonella O group N. Many O157:H7 strains may generate false negative reactions with H7 antisera and require several passages through motility medium for their accurate detection. In addition, atypical strains (O157:NM) will generate negative reactions due to the lack of the H7 antigen and hence require further biochemical testing. Recently developed PCR based methods for the detection of rfbE gene encoding O157 somatic antigen and fliC gene encoding H7 flagellar antigen can be used to detect both motile and non-motile strains of O157:H7 and offer a rapid, sensitive, specific, and more reliable alternative to LAT test (see below). 27.1.3.3 Enrichment and IMS Several factors influence the detection threshold of O157:H7 from clinical material, food, and cattle feces. In human patients, the recovery of O157:H7 declines from >90% during the first 6 days of illness to 33% for stools collected at a later date.71 The concentration at which E. coli O157 is shed in cattle feces varies from 102 to 105 CFU/g, but most adult cattle (61–85%) excrete less than 100 CFU of O157:H7 per gram of feces with high titers of background microflora.72–75 The numbers of O157:H7 can vary greatly from sample to

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sample within the same fecal pat.76,77 Studies have indicated that beef products may contain as little as 0.3–15 CFU of O157:H7 per gram78,79 and certain acidic foods, such as apple cider, may contain acid injured O157:H7 that require special recovery procedures.80 Consequently, selective enrichment followed by IMS of O157:H7 is often necessary prior to plating on selective media.75,77,81-83 Several protocols have been described for isolating O157:H7 from food, environmental, and clinical matrices.84 Variables such as the type of enrichment broth (trypticase soy broth; TSB, E. coli broth; EC or buffered peptone water; BPW), the selective agents (bile salts and antibiotics such as cefixime, cefsulodine, and vancomycin), the incubation temperature (35–37ºC), and the duration of incubation period (6–24 h) may influence the sensitivity of detection from different sample matrices.85 The comparison of efficacy of different enrichment protocols used by different authors was recently reviewed by Vimont et al.;84 the general consensus is that, compared to direct plating on selective media, enrichment procedures improve detection threshold of O157:H7 by 10- to 100-fold. For instance, direct plating of cattle fecal samples on CT-SMAC was compared to enrichment in trypticase soy broth containing selective antibiotics followed by plating on CT-SMAC. In this case, enrichment increased the detection limit from 251 CFU/g by direct plating to 13–16 CFU/g by enrichment.86 IMS has greatly improved the detection sensitivity of O157:H7 from enrichment cultures of different matrices. IMS uses commercially available magnetic beads coated with antibody against E. coli O157 antigen (e.g., Dynabeads anti-E. coli O157; Dynal, Inc., Lake Success, N.Y.) for separation and concentration of O157:H7 from mixed cultures. Conventional IMS procedure consist of mixing a sample (usually enriched cultures) with antibody coated paramagnetic beads in a microcentrifuge tube. Once the target organism (O157:H7) is captured by the antibody coated beads, a magnet is applied to the sides of the tube to hold magnetic beads and the liquid sample that contains most of the background flora is removed from the tube. The beads are then washed to remove the additional background material and resuspended in a small volume of buffer followed by plating onto selective or differential media. Several studies have shown the usefulness of enrichment and IMS methods for a range of clinical, food, and environmental samples.2,3 For instance, a study that compared direct plating of bovine feces on CT-SMAC with that of enrichment and IMS procedures prior to plating on CT-SMAC showed that this combination of procedures improved the detection sensitivity of O157:H7 by 10- to 100-fold as compared to direct plating of bovine feces.75 Several modifications of the standard IMS procedure have been made, including modifications in apparatus, sample volume, bead sizes, and washing procedures.87–90 Although, conventional IMS procedures are not technically complex, the process is labor intensive and not amenable to high sample throughput. Consequently, an automated IMS method with an integrated ELISA (EiaFoss, Foss Electric A/S, Hillerod, Denmark) has also been developed that

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allows rapid automated sample processing. The EiaFoss automatically carries out an immunomagnetic concentration of E. coli O157:H7 organisms and subsequently carries out an ELISA. Twenty-seven samples can be processed during one run, with an analysis time of about 100 min. Reinders et al.88 compared the sensitivity of an automated EiaFoss IMS procedure to that of conventional IMS for the detection of O157:H7 from milk. They reported that both methods were equally sensitive, but automated IMS provided a rapid and efficient alternative for manual IMS procedure. A recent modification of IMS procedure includes the use of an intrasolution magnetic particle separation device, called PickPen (BioNobile, Turku, Finland). The PickPen is available as the eight channel device that enables higher throughput processing of samples in 96-well format and offers a significant advantage over the single-tube IMS procedure. Nou et al.91 evaluated the usefulness of PickPen IMS for the detection of O157:H7 from enrichment cultures of cattle feces, hides, carcasses, and ground beef and compared this procedure with conventional IMS. They reported that PickPen IMS was significantly more sensitive than conventional IMS and that the PickPen IMS procedure greatly increased the throughput of IMS testing. Although enrichment and IMS procedures have improved the detection sensitivity of conventional culture procedures, these procedures are tedious and take an additional 16–24 h prior to plating onto selective media. Hence, a combination of pre-enrichment, IMS, and plating on selective media followed by screening of suspect O157:H7 colonies by conventional biochemical and serological testing may take several days (about a week) before the confirmatory detection of O157:H7 can be achieved. Recent studies have shown that the alternative approaches that combine molecular techniques such as PCR and microarrays with that of selective enrichment and IMS procedures can reduce the time required for confirmatory detection of O157:H7, and can also improve the detection sensitivity and specificity. 27.1.3.4 ELISA During the last two decades, several immunological methods (e.g., ELISA/EIA, colony blot, passive agglutination assays) have been developed for the detection of O157:H7 from multiple sample matrices. A number of these immunological assays are now available as commercial “ready to use test kits” for routine testing in clinical as well as food microbiological laboratory settings. These tests detect either the presence of Shiga toxins in stool samples and culture supernatants or detect bacterial antigens such as O157 and H7 from stools, food, and environmental samples. A commercially available Premier EHEC (Meridian Bioscience, Inc., Cincinnati, OH) ELISA is designed to detect Stx1 and Stx2 from diarrheal stool specimens or broth cultures of clinical isolates. This is a rapid 96-microwell enzyme immunoassay that uses monoclonal anti-Shiga toxin antibody absorbed to microwells. Addition of supernatant solution from test samples to the microwells permits binding of Stx to the monoclonal antibodies. This step is followed

Molecular Detection of Foodborne Pathogens

by the addition of an enzyme conjugated anti-Stx polyclonal antibody and a substrate for the colorimetric detection of Stx toxin. The Premier EHEC test has been evaluated by several researchers for the detection of Shiga toxins produced by E. coli including O157:H7 from clinical specimens.92–99 Most of these studies reported good sensitivity and specificity of the test, although one limitation at the microbial community level is that it is not possible to link presence of Stx toxin to any strain or serovar. Thus, this test at best provides rapid tool for screening of clinical specimens for the presence of Stx producing E. coli (including O157:H7) and further confirmation of O157:H7 by microbiological testing is often required. Although Premier EHEC test is a good presumptive test for Stx producing E. coli from clinical stool specimens, Hyatt et al.100 evaluated its performance on bovine feces and reported that this test had a poor sensitivity for detection of O157:H7 from these samples. It has also been reported that this test can produce false positive results with Pseudomonas aeruginosa isolates101 and may require overnight incubation of some strains to detect these toxins.102 The ability of Premier EHEC test to detect variants of Shiga toxins other than Stx1, Stx2, and Stx2c has not been explored. Another commercially available test is ProspecT Shiga toxin (STEC) microplate assay (Lexon-Trend, REMEL, Lenexa, KS). This assay is similar to the Premier EHEC except that the microtiter wells are coated with a polyclonal anti-Stx1 and Stx2 antibody and the secondary antibody is a horse radish peroxidase conjugated monoclonal anti-Stx1 and Stx2 antibody. Gavin et al.103 evaluated the performance of ProspecT assay for the detection of Shiga toxin producing E. coli in stool samples and reported that this assay was 100% sensitive and specific for detection of E. coli O157 in stools compared with SMAC. In addition, the ProspecT assay detected twice as many STEC as SMAC. This test has not been validated for detection of all known variants of Stx1 and Stx2. The Ridascreen assay (R-biopharm, Darmstadt, Germany) is also commercially available for detection of Stx toxins produced by STEC isolated from various sources. Bonardi et al.104 compared the performance of Ridascreen assay with that of vero cell test, PCR, and EIA ELISA using 34 strains of O157:H7 strains isolated from bovine feces and carcasses. These authors reported that Ridascreen assay was as sensitive as PCR, with less time required and simpler execution. This assay was also used successfully to investigate a multistate outbreak of E. coli O26:H11 in Germany.105 Recently, the Ridascreen assay was reported to detect strains producing Stx1 and variants Stx1c and Stx1d, as well as Stx2 and variants Stx2d1, Stx2d2, Stx2e, Stx2d, Stx2-O118 (Stx2d-ount), Stx2-NV206, Stx2f, and Stx2g.106 This assay showed a relative sensitivity of 95.7% and a relative specificity of 98.7%. Some of the Stx2-O118, Stx2e, and Stx2g producing STEC were not detected. Although the above immunoassays are not specific for the detection of O157:H7, they provide a straightforward approach for the presumptive identification of virtually all Shiga toxin producing E. coli (including O157:H7) in clinical samples. From a clinical perspective, administration of potentially deleterious

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antimicrobial or antimotility treatments would be avoided due to the rapid detection of STEC in patients suffering from HC and HUS.107 Rapid immunological assays for the detection of O157 and/or H7 antigens have also been developed. One such assay is O157 ELISA (LMD Laboratories, Inc., Carlsbad, CA) that uses microwell test strips coated with anti-E. coli O157 polyclonal antibody to detect O157 antigen from clinical samples. Dylla et al.108 and Park et al.109,110 compared this O157 ELISA assay with culture for the detection of E. coli O157:H7 from human stool samples and reported that, as compared to SMAC plating, this assay was a highly sensitive and specific method for screening stool samples for O157:H7 and required <1 h for completion. This method overcomes the limitation of SMAC plating because sorbitol fermenting O157 can also be detected. Nevertheless, the assay could not differentiate toxigenic from non-toxigenic strains and culture isolates of Citrobacter freundii, Esherichia hermanii, and Salmonella urbana produced false positive ELISA results. Several other test kits [e.g., Now-E. coli (Binax, Portland, ME); Path-Stik O157 (lumac, Landgraaf, Netherlands); VIP (BioControl system, Bellevue, WA); EHEK-Tek ELISA system (Organon Teknika, Durham, NC); Rapiblot E. coli O157 (Kalys Bioscience, Nepean, Ontario, Canada); ImmunoCard STAT-E. coli O157:H7 (Meridian Diagnostics, Inc., Cincinnati, OH); Reveal (Neogen, Lansing, MI)] are also commercially available and several performance evaluations have been published. For instance, in a multicenter prospective and retrospective study, the performance of a rapid ImmunoCard STAT-E. coli O157:H7 was evaluated for the detection of O157:H7 in stool samples.111 Fourteen (100%) specimens positive by culture for O157:H7 were found positive by the ImmunoCard STAT O157:H7 test, and there were no false positives from 263 culture-negative specimens. In a retrospective study, the test was positive in 339 (81%) of 417 stored culture-positive specimens and the specificity was 95% (98 of 103 specimens) with no false positives associated with alternate stool pathogens.111 Onoue et al.112 conducted a multicenter study to evaluate the performance of various plating methods with immunological methods including Now E. coli, Path-Stik O157, VIP, EHEC-Tek ELISA system and Rapiblot E. coli for the detection of O157:H7 from spiked radish sprouts and ground beef samples. They reported that all the immunological methods were as sensitive as plating methods, but their specificity was lower because false positives were obtained from uninoculated samples. Chapman and Ashton113 compared the efficacy of IMS-plating with that of Reveal, STAT, and VIP for the detection of O157:H7 in enrichment cultures of artificially spiked beef carcasses. They reported that IMS-plating method detected O157:H7 from 32 out of 32 meat samples spiked with an initial inoculum of 10 CFU/g, but only 30, 19, and 9 samples were positive by Reveal, VIP, and STAT, respectively. This study indicated that although these immunoassays were simple and rapid to use, the sensitivity of these assays fell short as compared to the sensitivity of the culture and this would limit their usefulness for routine screening of food samples.

Recently, several immunoassays (E. coli Now, Reveal, and VIP) were compared with IMS methods for the detection of O157:H7 from artificially spiked and naturally contaminated fecal, hide, carcass, and ground beef samples.114 They reported varying sensitivities of ELISA assays for different sample types in which, when detection results of ELISAs were compared with IMS, poor agreement was observed for fecal samples (kappa=0.10, 0.02 and 0.03 for E. coli Now, Reveal, and VIP, respectively), and fair to moderate agreement was observed for hide samples (kappa=0.30, 0.51, and 0.29 for E. coli Now, Reveal, and VIP, respectively), but there was near to perfect agreement between IMS and ELISAs for ground beef samples (kappa=1, 1, and 0.80 for E. coli Now, Reveal, and VIP, respectively). In addition to these, several studies have compared commercial as well as custom made ELISAs to traditional culture methods or IMS for the detection of E. coli O157:H7 from various food samples such as ground beef, lettuce, lamb products, beef carcass samples as well as bovine feces.114–121 In general, studies of comparative specificity and sensitivities of ELISA assays indicate that application of these assays to detect either Stx toxins or to detect O157 and/or H7 antigens is likely to be very useful for clinical and food microbiology laboratories. Detection of Stx toxin in particular allows detection of several Stx producing E. coli and therefore facilitate broader screening for several potentially pathogenic E. coli. Nevertheless, due to the potential for false positive or false negative ELISA reactions and lower sensitivity in some cases, independent confirmation of isolates by PCR based identification of virulence genes (e.g., stx1, stx2, eaeA) and serotype specific genes (rfbE and fliC) or by isolation and microbiological identification of O157:H7 would be required to facilitate confirmatory detection of O157:H7.

27.1.4 Molecular Diagnostic Techniques 27.1.4.1  PCR In recent years, PCR has been used increasingly both to confirm the identity of E. coli O157:H7 isolates and for rapid, sensitive, and serotype-specific detection of low levels of O157:H7 in feces, foods, water, and environmental samples. Several virulence associated as well as serotype-specific genes such as rfbE that encodes O157 lipopolysaccharide O-antigen,122 fliC encoding H-7 specific flagellar antigen,123 eaeA encoding intimin,51,124 hlyA encoding hemolysin,125 uidA (gusA) encoding β-glucuronidase,126 stx1 and stx2 encoding Stx1 and Stx2 toxins, respectively,127 have been used for genotyping and detection of O157:H7 in both standard and multiplex PCR (m-PCR) formats. One limitation of a single gene PCR assays is that there are no single markers that provide definitive identification for O157:H7. This is primarily because E. coli O157:H7 is genetically similar to most other E. coli strains that are commonly found in gastrointestinal tract of human and animals and most of the virulence associated genes targeted for PCR amplification are widespread in non-O157 E. coli isolates. For instance, genes encoding Stx are widespread in several non-O157

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STEC isolates and Stx negative strains of O157:H7 have also been reported.128 Similarly, the eaeA gene is widespread in serogroups belonging to enteropathogenic E. coli (EPEC).2 The hlyA gene is present in several non-O157 serogroups125 whereas the rfbE gene is also found in all isolates of E. coli belonging to O157 serogroup, and fliC gene is found in all serotypes belonging to H7 serogroup. Therefore, single gene PCR as well as multi-gene PCR assay, that do not incorporate a proper combination of both virulence markers (e.g., eaeA, stx1, and stx2) and serotype-specific markers (e.g., rfbE and fliC) often lack specificity and require further confirmation using conventional biochemical typing. In the mid-1990s, Fratamico et al.129 developed an m-PCR assay based on detection of eaeA, stx, and ehx genes from O157:H7 isolates. This PCR assay detected all three genes from O157:H7 isolates, but the assay lacked specificity because none of the genetic markers were specific to O157:H7 and hence was prone to generate false positive results. Another m-PCR targeting a highly conserved O157:H7 serotype-specific allelic substitution in uidA gene in combination with primers amplifying stx1 and stx2 genes was shown to be specific for the detection of both O157:H7 and its non-motile variant, O157:NM.130 Noteworthy is that most E. coli strains including most pathogenic strains (except O157:H7), produce functional β-glucuronidase enzyme encoded by uidA, but all O157:H7 strains carry the entire uidA gene in its chromosome with several base mutations out of which +93 T to G mutation is highly conserved among both typical and atypical strains of O157:H7;126 thus, this mutation can be used as a target for the PCR based identification of O157:H7. Although the above PCR method was specific for typical strains of O157:H7, it did not include the eaeA marker. Detection of both eaeA and stx genes is important because isolates (especially atypical strains) may lose stx genes upon subculture and stx negative strains of O157:H7 have also been isolated.128,131 Meng et al.132 developed m-PCR assay to simultaneously detect three virulence markers (eaeA, stx1, and stx2) from several O157:H7 isolates, however none of the serotype-specific markers (rfbE and fliC) were incorporated into their assay. To improve the specificity, an m-PCR assay was developed in which primers targeting H7 flagellar antigen gene, fliC, were combined with the primer sets that amplify stx and O157specific C′-terminal region of eaeA gene.133 Although both virulence genes and a serotype-specific H7-flagellar antigen gene were combined, the assay had pitfalls because the primers that amplify O157 polysaccharide (rfbE) gene were not included and the amplification product from H7-flagellar gene specific primers would generate false positive with other non-O157 serotypes.134 For the serotype-specific identification of O157:H7, the primer pairs that specifically amplify O157-specific rfbE gene and H7-specific fliC gene were combined in another m-PCR reaction,135 however this assay did not identify any of the virulence genes. Finally a robust m-PCR assay was developed that amplified a combination of fliC, rfbE, stx1, stx2, and eaeA genes that allowed serotypespecific detection of all pathogenic O157:H7 strains and their differentiation from non-O157 E. coli.136 The efficacy of this

Molecular Detection of Foodborne Pathogens

m-PCR assay for the detection of O157:H7 DNA extracted directly from artificially spiked cattle feces was compared with that of DNA extracted from TSB enrichment cultures.136 The authors found that the m-PCR assay could detect as few as 1 CFU/g of feces (initial inoculum) following TSB enrichment, but often failed to detect the organisms directly from fecal samples (without prior enrichment) when using a QIAmp test kit (Qiagen) to extract template DNA. The efficacy of this PCR assay was also evaluated for the detection of O157:H7 from water and soil samples spiked with different concentrations of the bacteria.137 Soil and water samples were subjected to 6 to 8 h enrichment prior to m-PCR. These researchers reported the detection limits of 1 CFU/ ml of drinking water and 2 CFU/g of soil. The starvation of O157:H7 for 35 days prior to addition to soil did not affect the sensitivity of the assay to detect initial cell numbers as low as 10 CFU/g of soil. Fratamico et al.138 developed the m-PCR using a combination of primers to co-amplify hlyA, fliC, stx1, stx2, and eaeA genes of O157:H7 and applied this PCR assay on DNA extracted from 24 h enrichment cultures of ground beef, blue cheese, mussels, alfalfa sprouts, and bovine feces. All samples were artificially inoculated with various levels of O157:H7 strain 933. This assay had a detection limit of ≤1 CFU/g (initial inoculum level) with results obtained within 24 h. The limitation of this assay was that it did not include the primers to amplify O157-specific rfbE gene and hence may still require further confirmation if the assay is to be used for routine screening of samples. The success of any PCR based pathogen screening method is directly dependent on the efficacy of nucleic acid extraction method employed. Stewart et al.139 evaluated several procedures including boiling, enzyme treatment, enzyme treatment plus phenol chloroform isoamylalcohol (PCI) extraction, and enzyme treatment followed by PCI plus Geneclean (Bio 101, Vista, CA) purification, for preparing template DNA from unenriched and enriched cultures originating from cattle feces. Using PCR based detection of stx1 gene they found that the boiling method of template DNA was most sensitive as it allowed detection of as few as 3 CFU/g (initial inoculum) from enriched fecal cultures and 105 CFU/g from nonenriched cultures. When the boiling method was combined with IMS, the detection sensitivity of their PCR assay was only 103 CFU/g of feces. McKillip et al.140 used pre-treatment of various dairy food samples (skim milk, nonfat dry milk, cheddar, brie cheese and reconstituted whey powder) with a combination of petroleum ether, ethanol, and ammonium hydroxide prior to DNA extraction using two methods (i) PCI extraction, or (ii) bacterial concentration followed by guanidinium isothiocyanate treatment and PCI extraction. PCR detection limits after each DNA recovery method varied with the specific food, ranging from 10 to 104 CFU/ml for all dairy products except for whey powder. DNA yields and subsequent PCR detection limits for reconstituted whey powder were very poor, and neither procedural changes nor the addition of PCR enhancement agents were able to improve recovery and/or detection. Johnston et al.141 reported that DNA prepared from preconcentrated organisms, using

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high speed centrifugation followed by extraction using the Plant DNAzol kit (Invitrogen, USA), allowed sensitive and direct detection (without prior cultural enrichment) of stx and invA (S. typhimurium) genes from artificially spiked alfalfa sprouts and spent irrigation water (10 CFU/g and 10 CFU/ml, respectively). Tortorello et al.142 compared six methods [direct fluorescent antibody (DFA), antibody-direct epifluorescent filter technique (Ab-DEFT), direct selective plating on sorbitol MacConkey agar (SMA), IMS coupled to either selective plating (IMS-SMA) or the polymerase chain reaction (IMSPCR), and flow cytometry (FC)] for the detection of O157:H7 in enriched and nonenriched cultures of inoculated apple juice. The most sensitive detection, 0.1 CFU/ml of O157:H7, was provided by the IMS-SMA and IMS-PCR after 8 h of enrichment. Absolute detection limits (without enrichment) were 103 CFU/ml for IMS-PCR and 104 CFU/ml for IMSSMA. As few as 10 CFU (initial inoculum) of O157:H7 could be detected per ml of apple juice after 28 days of 4ºC storage followed by 8-h enrichment and IMS-PCR or IMS-SMA. Fitzmaurice et al.143 used a combination of a membrane surface adhesion technique and m-PCR assay to detect O157:H7 from 16 h enrichment cultures of minced beef spiked with O157:H7 and compared this method with IMS-PCR. The template DNA was extracted from polycarbonate membrane immersed in an enrichment culture and from IMS beads by enzymatic lysis followed by PCI extraction. These authors reported that similar numbers of organisms could be detected by both IMS-mPCR and membrane capture-mPCR when E. coli was present at high concentrations in enrichment samples (log10 9.6–7.5 CFU/ml). At levels lower than log10 6.4, however, only the IMS method could recover bacteria. In another study, the efficacy of IMS and PCR was compared for the detection of O157:H7 from 8 h enrichment cultures of animal feces and various meats spiked with O157:H7.144 While the IMS assay was ten times more sensitive than the PCR assay for most samples, the lowest level of the inoculum at which 100% of the replicate samples were found positive by PCR was 102 CFU/g (cattle and sheep feces), 10 CFU/g (raw mince beef, dry fermented sausage) or 1 CFU/g (fliet americain). These authors also confirmed the utility of the boiling method for DNA template preparation along with incorporation of 25 µg of BSA in a PCR reaction to overcome problems of PCR inhibitors. Visetsripong et al.145 tested the efficacy of m-PCR to detect stx1 and rfbE genes from enrichment cultures of raw meat spiked with varying concentrations of O157:H7. These authors reported the detection sensitivity of 102 CFU/ml when the template DNA was prepared by boiling of 8 h enrichment cultures followed by incorporation of 100 µg of BSA in a PCR reaction. The detection sensitivity improved to 10 CFU/ml when PCR was performed on 16 h enrichment cultures. Although PCR offers advantages such as rapid and specific detection of O157:H7, the application of PCR assays for the specific detection of O157:H7 directly from sample matrices such as feces or different types of foods is often challenging because these samples not only contain very low numbers

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of O157:H7 organisms (≤10–200 organisms), but also carry PCR inhibitory substances and a high amount of background flora. Moreover, PCR requires very small application volumes (1–10 µl) of DNA that is extracted from high sample volumes (≥25 ml or g) in the presence of matrix associated compounds that may interfere with enzymatic nucleic acid amplification. Therefore, the method of DNA preparation may have profound effects on the analytic sensitivity of any given assay. It appears that despite the exquisite sensitivity of PCR methods, successful application in real world matrices benefit greatly by including some type of sample enrichment. One advantage of including enrichment in the analytical process is that the practitioner is more likely to detect viable cells from these samples. 27.1.4.2 Real-Time PCR The presence of very low amounts of O157:H7 in various sample matrices, a very low infective dose of O157:H7 in humans (10–100 organisms),38 and difficulties in direct detection of O157:H7 from foods and other environmental samples have spurred development of alternative PCR assays. Realtime PCR (R-PCR) assays that incorporate the fluorogenic probe in a PCR reaction have been developed for the detection of O157:H7 in foods, feces, and environmental samples (Table  27.1). One of the most commonly used R-PCR systems is the TaqMan PCR detection system in which a fluorogenic oligonucleotide probe complementary to the target gene is labeled with a fluorescent reporter dye at 5′ end and a fluorescent quencher dye at the 3′ end. This probe binds to its complementary sequence within the target gene selected for amplification by flanking primers. During amplification, 5′→3′ nuclease activity of the Taq DNA polymerase cleaves the probe, separating the quenching dye from the reporter dye leading to the increased fluorescence emission. Fluorescent intensity is then measured by a fluorometer and the emission data is analyzed for the detection of a “positive” or “negative” result for the target genes; fluorescence intensity is proportional to copy number thereby allowing quantitative analyses. Incorporation of fluorogenic probe in the PCR assay offers capabilities of simultaneous detection as well as quantification of very low levels of target organisms in a sample with an additional element of specificity that eliminates the need for post-PCR analysis of PCR products by agarose gel electrophoresis. Moreover, because of the automation, these assays can be easily adapted for high-throughput applications. Hairpin structured fluorogenic nucleic acid probes, referred to as molecular beacons (MB), provide an alternative R-PCR detection system.146 The stem of MB is made of two short oligonucleotide arms that are complementary; one of the arms is terminally labeled with reporter dye and the other arm is labeled with a quencher dye. MB posses differential fluorescent properties based on the relative stability between its two duplex forms (hairpin and the probe target hybrid). In the hairpin conformation, the fluorescent reporter is immediately adjacent to the quencher so that the intensity of the background signal is minimized. Hybridization of the probe to single stranded target causes hairpin to open with the

378

Molecular Detection of Foodborne Pathogens

Table 27.1 Real-Time PCR Assays used for the Detection of O157:H7 from Various Sample Matrices Real-Time PCR

Target Gene

Fluorogenic TaqMan assay

stx1

Fluorogenic TaqMan assay

Fluorogenic TaqMan assay

eaeA

eaeA

Growth/Enrichment Conditionsa

DNA Preparation Method

Detection Limit (cfu)

Beef

mECn20 (37ºC/12 h)

Boiling

0.5/gb

Beef

None

Boiling

>5 × 103/gb

Pure culture

TSB (37ºC/ OD600=0.8)

Boiling

4 × 103/ml

Ground beef

mTSBn100, c10, v8, cf0.05 (37ºC/6 h)

QIAamp tissue kit (Qiagen) or DNA-ER solution (Perkin-Elmer)

≥104/gc

Pure culture

mTSBn100, c10, v8, cf0.05 (37ºC/4–8 h) or (followed by IMS) mECn20 (37ºC/4–8 h)   or (followed by IMS)

Sample

Beef

mTSBn20 (37ºC/6 h)

Cattle feces Pure culture

mTSBn20 (37ºC/6 h) TSB (37ºC/18 h)

Reference 148

150

≥103/ml or (102/ml)c

≥103/ml or (102/ml)c InstaMatrix (BioRad)+boiling

3/gb

151

1.2/gb 2 × 104/ml

Multiplex SYBR Green I assay

stx1 and stx2

Pure culture

NA

DNeasy mini kit (Qiagen)

1 cfu

193

Molecular beacon assay

rfbE

Apple juice

mECn20 (37ºC/11 h)

InstaMatrix (BioRad)+boiling

1/mlb

155

Milk Pure culture

mECn20 (37ºC/6h) mEC (37ºC/overnight)

Pure culture

NA

Human feces

None

Feces

mTSB (37ºC/16 h)

Ground beef Pure culture

mTSB (37ºC/16 h) mTSB (37ºC/16 h)

Feces

mTSB (37ºC/16 h)

Ground beef

mTSB (37ºC/16 h)

Soil

mLBc10, v8, cf0.05 (37ºC/16 h)

Soil, feces, and waste water

None

Multiplex molecular beacon assay

Fluorogenic TaqMan assay

Multiplex TaqMan assay

Multiplex assay

stx1 and stx2

eaeA

stx1 and stx2

eaeA, stx1, and stx2

1/mlb 5 × 103/ml ID-DNA extraction kit (Infectio Diagnostics, Canada)

10–50 cfu

147

105/gb Lysis+boiling+Qia Quick PCR purification kit (Qiagen)

15/gb

194

1.5/gb 103/ml Lysis+boiling+QIA Quick PCR purification kit (Qiagen)

1.2–12/gb

1.2/gb UltraClean soil, fecal, and water DNA kits (MO BIO, Inc., Solana Beach, CA)

1–10/gb

156

≥3.5 × 104/gb

Multiplex fluorogenic assay

stx1 and stx2

Pure culture

NA

Lysis+boiling

5 × 102 cfu

149

Multiplex assay

eaeA and hlyA

Pure culture

NA

Lysis+boiling

5 × 102 cfu

149

Multiplex TaqMan assay

eaeA, stx1, and stx2

Cattle feces

GN+TSB (37ºC/16 h)

Lysis+boiling+Qia Quick PCR purification kit (Qiagen)

2.5–25/gb

157

103/ml

Pure culture Fluorogenic assay

eaeA

Groung beef

IMS without enrichment

Lysis+heating

1.3 × 104/gb

152

Multiplex TaqMan assay

rfbE and stx2

Feces and apple juice

None

Boiling+QIAamp stool kit (Qiagen)

104 cfub

195 (Continued)

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Escherichia

Table 27.1  (Continued) Real-Time PCR

Target Gene

Sample

Growth/Enrichment Conditionsa

Beef and milk Milk Apple juice Pure culture

None TSB (37ºC/4 h) TSB (37ºC/10 h) TSB

DNA Preparation Method

Detection Limit (cfu)

Reference

103 cfub 10/mlb 10/mlb 103/ml

Fluorogenic TaqMan assay

eaeA

Sandy loam soil

None

FastPrep FP120 bead beating system (Bio-101, Vista, CA) followed by PCI

1.3 × 105/gb

153

Fluorogenic assay

eaeA

Ground meat

Two step enrichment in TSB (37ºC/2 h followed by 37ºC/3 h)

Membrane capture+PrepMan Ultra reagent (Applied biosystems, CA)

102/25gb

154

Fluorogenic TaqMan assay

eaeA/stx1/ stx2

Dairy wastewater

LB (37ºC/24 h)

AquaPure genomic DNA kit (Bio-Rad)

104/ml

158

a

b c

IMS, immunomagnetic separation; PCI, phenol chloroform isoamylalcohol; GN+TSB, Gram negative+trypticase soy broth; mEB, modified E. coli broth; mTSB, modified trypticase soy broth; mLB, modified luria broth; None, no growth or enrichment; NA, not applicable; v, vancomycin; c, cefsulodin; cf, cefixime; n, novobiocin; the numbers indicate concentration of each antibiotic (mg/ml). Detection limit is the actual numbers of bacteria present in the samples (either artificially spiked or natural sample) prior to enrichment. Detection limit is the actual numbers of bacteria present in the final enrichment sample.

result that quencher and reporter dye are no longer in close proximity of each other leading to emission of fluorescence. The use of MB in R-PCR offers supplementary level of specificity in which no increase in fluorescence is observed even with the presence of target strand that contains only a single nucleotide mismatch146. Most of the R-PCR assays that have been developed for the identification of O157:H7 are based on the detection of genes encoding Stx toxins,147–149 intimin,150–154 and O-antigen.155 These R-PCR assays offer the opportunity to quantify the absolute and relative amounts of O157:H7 in complex sample matrices (Table 27.1). Most of these assays target single genes, however, and therefore lack diagnostic specificity by themselves, which could lead to problems of cross reactivity (false positives) with other closely related organisms. For instance, R-PCR assays that target stx genes may cross react with other non-O157 STEC and fail to discriminate between O157:H7 and other E. coli. Some published reports have shown that the specificity and sensitivity of R-PCR assays can be improved by using a combination of two or more primer sets (multiplex-R-PCR assay) in one reaction156–158 although in practice is it not possible to multiplex more than four markers due to the limits of overlapping excitation of emission spectra of available fluorphores. Furthermore, several factors such as the initial numbers of O157:H7 in the sample, the type and volume of sample, presence of nontarget background microflora, PCR inhibitors and protocols used for the template DNA preparation may have significant influence on the sensitivity of the R-PCR assays used for the detection of O157:H7. As a result, the detection sensitivities of several published R-PCR assays for unenriched foods, feces, and environmental samples has been reported to be very low (>103–105 CFU; Table 27.1).

PCR inhibition is a common problem with DNA isolated from food, environmental, and clinical samples because of presence of substances such as heme in blood and meat samples and heavy metals and complex humic acids and fulvic substances in feces and soils, and polyphenolic compounds in acidic foods such as apple cider.155,159–164 Inhibitors can also squelch fluorescent signal from fluorophores used in these assays.165 The problems of PCR inhibitors can be partially dealt with further cleanup of DNA extracts and by spiking samples with internal PCR amplification controls such as those encoding green fluorescent protein (gfp) or targeting other sequences not present in O157:H7.147,166 Detection sensitivities of most R-PCR applications depend not only on the efficiency of nucleic acid extraction method, but can be improved by enrichment procedures that also increase likelihood of detection of viable cells. The enrichment step not only increases template numbers, but has the effect of diluting inhibitory substances. As with conventional PCR, the length of enrichment affects the detection sensitivity of R-PCR assays. Fortin et al.155 reported that when combined with selective enrichment step, R-PCR assay could detect as few as 1 CFU of O157:H7 organisms per ml in raw milk after 6 h of enrichment whereas for apple juice samples, the sensitivity of detection was much lower and a minimum of 11 h of enrichment was needed to achieve the same detection levels in raw milk. Similar results were obtained by Hu et al.136 who reported the sensitivity of the R-PCR assay to be 10 CFU/ml for milk samples and apple cider enriched for 4 h and 10 h, respectively. Sharma et al.151 reported that the sensitivity of R-PCR assay to detect O157:H7 from enriched cultures of beef samples was adversely affected by both the length of enrichment and the presence of non-target background microflora. They found that the detection sensitivity of

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R-PCR assay improved from 3 × 103 CFU/ml in 2 h enriched cultures to 3 CFU/ml when cultures were enriched for 6–18 h prior to PCR analysis even when beef samples harbored 103 CFU/ml of background flora. When background flora increased to 108 CFU/ml, the detection sensitivity of R-PCR assay was reduced by 100-fold. 27.1.4.3 Reverse Transcriptase PCR Reverse transcriptase (RT)-based assays have been developed for the detection of viable O157:H7 in foods and environmental samples. Conventional PCR and R-PCR assays rely upon a DNA template for amplification. This DNA can, theoretically, persist in the sample even after cell death thereby leading to positive detection of nonviable cells.167 This could be a significant disadvantage and could be construed as false detection events because nonviable bacteria present no health risks.168 Thus, detection of O157:H7 from food and environmental samples by conventional or R-PCR is generally followed by isolation and confirmatory identification of the pathogen with conventional procedures. An alternative approach relies on RT to convert mRNA templates to cDNA for subsequent PCR amplification. mRNA is present only in actively growing cells and unlike DNA, mRNA tends to degrade rapidly (half-life of mRNA varies from 40 s to 20 min) in dead cells, thus allowing detection of only viable cells in a sample.169 Yaron and Matthews168 tested the utility of RT-PCR for detecting several genes (rfbE, fliC, stx1, stx2, mobA, eaeA, hlyC, hlyA, and 16S rRNA) as an indicator of viability of O157:H7. They found that 16S rRNA, and mRNA of rfbE, stx1, and hly were detected in cells harvested during all growth phases whereas other mRNAs including stx2 were variable in different growth phases. The authors suggested that rfbE and stx1 mRNAs were more suitable targets for detecting both viable as well as nonculturable cells because these mRNAs were more stable in the viable cells, while having a very short half-life in the heat killed cells. They also reported that at least 107 CFU of O157:H7 are required if the RT-PCR is to be used to detect viable cells without enrichment, and that the conventional PCR was more sensitive than RT-PCR. McIngvale et al.170 showed that the ability of RT-PCR to detect stx2 mRNA from viable O157:H7 cells was influenced by growth conditions with the best detection of stx2 mRNA found with late log- and early stationaryphase cells incubated at 37ºC compared with cells incubated at 32ºC. They also reported that with an initial inoculum of 1 CFU/g of beef samples, detection of stx2 mRNA was possible after 12 h of enrichment. RNA extraction from heat killed cells produced no bands from RT-PCR, while PCR of stx2 DNA consistently produced bands following cell death. While the RT-PCR assays targeting rfbE,168 stx2170 or a combination of stx1 and stx2171 are suitable for the rapid and sensitive detection of viable O157:H7 cells, these assays can produce false positive results for those E. coli strains that harbor either stx1 or stx2 or that possess O157-type O antigen. As a result, Sharma et al.172 developed a RT-realtime multiplex RT-PCR assay (rRT-mPCR) targeting mRNA encoded by rfbE and eaeA genes for the specific detection of

Molecular Detection of Foodborne Pathogens

O157:H7 in a single step. This assay allowed specific detection of O157:H7 at a concentration of as low as 1 CFU/g of bovine feces (initial inoculum) using RNA prepared from 5 h enrichment cultures of artificially seeded bovine feces. The only pitfall of this assay was that it did not include the stx1 and stx2 alleles and therefore would require further confirmation. Another rRT-mPCR assay was designed to detect and identify O157:H7 using three combinations of primers and probes targeting eaeA, rfbE, and stx2 mRNAs.173 Although this assay did not include primers to amplify stx1 mRNA, these authors reported that rRT-mPCR assay was effective in differentiating E. coli O157:H7, non-O157 STEC and non-E. coli pathogens from 100 strains isolated from clinical patients and the environment. They also reported that the sensitivity of rRT-mPCR for detection of O157:H7 from various foods (milk, alfalfa sprots, and ground beef), feces and pure cultures without enrichment or prior treatment ranged from 103 to 106 CFU. When combined with enrichment and IMS procedure, the assay could detect as few as 10 CFU/ ml (initial inoculum) of O157:H7 from pure culture, foods, and stool. Finally, a sensitive RT-PCR method for specific detection of viable O157:H7 cells, including viable but nonculturable (VBNC) cells, in water samples was developed by Liu et al.174 The VNBC cells were generated through starvation of O157:H7 in deionized water and confirmed by culture and viability staining using Live/Dead BacLight bacterium viability kit (Molecular Probes, Europe, Leiden, The Netherlands). This method involved capture of the bacterial cells on a low-protein-binding membrane and direct extraction and purification of RNA followed by RT-PCR and electronic microarray detection of rfbE and fliC from O157:H7. This assay was reported to detect as few as 1 CFU of O157:H7 in diluted cultures, 3 to 4 CFU/liter in tap water, 7 CFU/liter in river water, and 50 VBNC cells in 1 liter of river water, demonstrating the best limit of detection reported to date for VBNC cells in environmental water samples. The success of RT-PCR requires that the target mRNA molecule has a short half-life and that the target mRNA is constitutively expressed in viable cells, including healthy, dormant, injured or otherwise nonculturable cells. Furthermore, the expression of bacterial virulence factors is often influenced by various environmental and stress conditions that organisms may encounter in different food matrices and environments. For instance Stx production has been shown to be reduced with reduced aeration and reduced growth temperature (32ºC).170,175 Therefore, to use RT-PCR as a means of specifically identifying bacteria, care must be taken to ensure abundant and dependable expression of target mRNA. 27.1.4.4  Microarray Nucleic acid microarrays have also been employed for detection and genotyping of bacterial pathogens.176,177 Microarrays are well known for their application in the whole genome expression profiling of bacterial pathogens and several reviews of gene expression studies have been published.178–180 From food safety perspective, microarrays can also be used for bacterial gene identification as well as pathogen

381

Escherichia

surveillance.165,181–185 For these applications, labeled nucleic acid targets (PCR products, genomic DNA, rRNA, etc.) are hybridized to a microarray chip whereupon target:probe duplexes are typically detected using some type of direct or indirect fluorescent signal system. The relative signal from each probe (also called a spot or feature) is detected and quantified by using a confocal laser or filtered light scanner. In this way, the presence or absence of specific gene can be detected. The major advantage of the microarray assay over agarose gel analysis of the PCR products is that microarray do not rely on the length of the PCR products for identification, but require that the internal sequences of the target genes to be complementary to the probes spotted onto the chip. Such hybridization methods are generally sensitive to sequences mismatches >10% from the target sequence depending on the type of probe and position of the mismatch sequences. While microarrays are commonly coupled with m-PCR assays, there are situations where they can be used independent of target amplification and thereby avoid limitations in the number of genes that can be detected. Furthermore, when used to detect sequence polymorphisms in sequences that are “universally” present (e.g., 16S rDNA), microarrays can simultaneously distinguish between many different bacterial pathogens.186,187 As a proof of concept Chizhikov et al.183 constructed a microarray chip to detect six virulence genes common to Shigella spp., O157:H7 and enterovirulent E. coli. The presence or absence pattern of the six virulence genes (eaeA, stx1, stx2, fliC, rfbE, and ipaH) was used to test unidentified isolates. Fluorescent dyes were directly incorporated into products via m-PCR assay and hybridized to 25 bp single-stranded oligonucleotides spotted on a glass slide. The authors reported that the assay was relatively fast, flexible, and more reliable for detecting genetic markers as compared with m-PCR alone.183 Jin et al.184 designed a multi-pathogen detection microarray by incorporating multiple genes for the simultaneous detection of O157:H7 (stx1, stx2, and uidA genes) and Vibrio cholerae (ctxA, tcpA, and LPSgt genes) isolates. Using a recombinant plasmid and target pathogens, they reported bench top analytical sensitivities of 102 copies and 103 CFU/ml per reaction, respectively. For the samples with fewer O157:H7, culture enrichment prior was required to avoid false negatives. Call et al.181 designed a nucleic acid microarray composed of 25–30-mer oligonucleotide probes complementary to four target gene sequences (eaeA, stx1, stx2, and hlyA). Target DNA was amplified by m-PCR using biotinylated primers and PCR products were hybridized to the array without further modification or purification. When coupled with enzymatic amplification the array was reportedly 32-fold more sensitive than gel electrophoresis and the benchtop analytical sensitivity of the assay was <1 cell equivalent of gDNA (1 fg). Furthermore, the combination of immunomagnatic capture, PCR, and a microarray analysis allowed detection of as low as 55 CFU/ml of a chicken carcass rinsate without the need for pre-enrichment. This represents an advance over PCR approaches that require enrichment as it reduces time required for analysis while

retaining high sensitivity. Chandler et al.165 coupled an electromagnetic fluidic cell system for the automated IMS of O157:H7 from poultry carcass rinsate. In this case, porous nickel foam was used to enhance the magnetic field within the flow path to ensure immobilization of the immunomagnetic particles throughout the fluid. O157:H7 cells were reproducibly isolated directly (without enrichment) from poultry rinsate with 39% recovery efficiency at 103 CFU/ ml inoculum. When recovered beads were used for direct PCR amplification and microarray detection, the detection limit of <103 CFU/ml was achieved. Recently, Jin et al.182 designed a microarray chip that included four O157:H7 gene targets (rfbE, fliC, stx1, and stx2) and reported that microarray chip was more sensitive than PCR alone and could be used for the specific and rapid detection of all tested O157:H7 isolates. Thus, when coupled with PCR, microarray based analysis can improve analytic sensitivity, diagnostic sensitivity, and diagnostic specificity of O157:H7 and these methods offer attractive advantages over conventional detection systems. Nevertheless, microarray-based assays are limited by the requirement for expensive instrumentation and the reagents used for microarray assays may be prohibitively expensive in the context of surveillance programs. Furthermore, sample throughput is constrained significantly when glass-slide formats are used.176 Some of the technical problems associated with performance and data interpretation of a microarray include high background interference (noise) especially at low signal output, and the differences in efficiency of nucleic acid labeling and variations in the experimental conditions in each laboratory can affect the interpretation of results. Care is also needed in design probes as secondary and tertiary structure can inhibit probe:target binding.188 Considerably more work is needed for inter-laboratory validation of microarray-based assays before this technology is likely to impact our ability to routinely test for O157:H7.

27.2  Methods It is important to recognize that while there are many published protocols for the molecular detection of O157:H7 and presumably some of these are in use, there is no single protocol that is widely viewed as “best” especially for the direct detection of O157:H7 from diverse matrices including foods, feces, and environmental samples. The protocol that is described here is a combination of US-FDA method for enrichment and isolation of E. coli O157:H7 from foods189 followed by definitive detection of presumptive O157:H7 colonies by a m-PCR136 that has been extensively validated by various researchers for the detection of virulence markers (stx1, stx2, and eaeA) and serotype-specific markers (fliC and rfbE) of O157:H7 isolated from a wide variety of sample matrices.77,137,190–192 It is expected that all O157:H7 colonies will generate either of the following profile of PCR amolicons: (i) 210 bp (stx1), 484 bp (stx2), 368 bp (eaeA), 292 bp (rfbE), and 625 bp (fliC); (ii) 484 bp (stx2), 368 bp (eaeA),

382

292 bp (rfbE), and 625 bp (fliC) or (iii) 210 bp (stx1), 368 bp (eaeA), 292 bp (rfbE), and 625 bp (fliC).

27.2.1 Reagents and Equipment (1) Balance, sensitivity of 0.1 g. (2) Stomacher 400 or 3500 with appropriate sizes of sterile Stomacher bags, with or without mesh or equivalent bag mixer and bags. (3) Incubator 37 ± 0.5°C. (4) Micropipettors to deliver 15–1000 μl with sterile disposable filtered micropipet tips. (5) Mechanical Pipettor with 1 ml, 5 ml, and 10 ml sterile pipettes. (6) Inoculating loops and spreaders. (7) Microcentrifuge and sterile 1.5 ml, microcentrifuge tubes. (8) PCR tubes and PCR thermal cycler (iCycler, BioRad, USA or equivalent). (9) Horizontal gel electrophoresis apparatus and DC power supply. (10) EHEC enrichment broth (EEB): trypticase soy broth (30 g); bile salts No. 3 (1.5 g); dipotassium phosphate (1.5 g) and deionized water to make 1 lit. Supplement with cefixime (0.0125 mg, Dynal inc., Lake Success, NY); cefsulodin (10 mg); vancomycin (8 mg). (11) CT-SMAC agar: Peptone (20 g); sodium chloride (5 g); bile salts No. 3 (1.5 g) ; sorbitol (10 g); crystal violet (0.001 g); neutral red (0.03 g); agar-agar (15 g) and deionized water to make 1 l. Supplement with cefixime (0.05 mg) and potassium tellurite (2.5 mg, Dynal Inc., Lake Success, NY). (12) TBE electrophoresis buffer (1 ×): Tris base (89 mM); boric acid (89 mM); EDTA (2 mM) and deionized water to make appropriate volume. (13) Electrophoresis-grade agarose. (14) 10× Loading buffer: Bromophenol blue (0.25 mg); xylene cyanol (0.25 mg); glycerol (25 ml) and deionized water to make 50 ml. (15) Ethidium bromide (10 mg/ml).

27.2.2 Sample Preparation (1) Weigh 25 g of food into 225 ml of EEB, blend or stomach briefly as necessary. (2) Incubate at 42 ± 0.5°C with shaking for 18–24 h (see notes i and ii). (3) Streak 0.1 ml of enrichment broth on CT-SMAC and incubate at 37°C for 18–24 h. (4) Pick up to 2 - 5 sorbitol negative colonies (see note iii) from CT-SMAC (per sample) and transfer to microfuge tube containing 200 µl of sterile deionized water. (5) Boil for 10 min and centrifuge at maximum speed for 10 min. Now the samples are ready for PCR analysis (see note iv).

Molecular Detection of Foodborne Pathogens

27.2.3  Detection Procedure (1) Use following primer sequences for the m-PCR assay.

Primer FliCh7-F FliCh7-R Int-F Int-R Rfb-F Rfb-R Slt-IF Slt-IR Slt-IIF Slt-IIR

Product Target Size (bp) H7

625

Intimin

292

O157

368

Stx1

210

Stx2

484

Sequence (5′–3′) GCGCTGTCGAGTTCTATCGAGC CAACGGTGACTTATCGCCATTCC GACTGTCGATGCATCAGGCAAAG TTGGAGTATTAACATTAACCCCAGG GTGTCCATTTATACGGACATC CATG CCTATAACGTCATGCCAATATTGCC TGTAACTGGAAAGGTGGAGTATAC GCTATTCTGAGTCAACGAAAAATAAC GTTTTTCTTCGGTATCCTATTCCG GATGCATCTCTGGTCATTGTATTAC

(2) Prepare following PCR reaction mixture in a 0.2-ml PCR tube. Component (concentration)

Final Concentration

Volume (µl)

Buffer (10×)

10 mM Tris-HCl (pH 8.3), 50 mM KCl, 0.1% gelatin, 0.1% Tween 20, 0.1% Nonidet P-40 2.5 mM 200 µM 60 nM 60 nM 75 nM 75 nM 100 nM 100 nM 200 nM 200 nM 200 nM 200 nM 2.5 U -

10

MgCl2 (25 mM) dNTPs (10 mM) Primer FliCh7-F (10 µM) Primer FliCh7-R (10 µM) Primer Int-F (10 µM) Primer Int-R (10 µM) Primer Rfb-F (10 µM) Primer Rfb-R (10 µM) Primer Slt-IF (10 µM) Primer Slt-IR (10 µM) Primer Slt-IIF (10 µM) Primer Slt-IIR (10 µM) Taq DNA polymerase (5 U/µl) Template DNA Deionized water Total

10 2 0.6 0.6 0.75 0.75 1 1 2 2 2 2 0.5 5–20 To make 100 µl 100

(3) Mix reagents and perform PCR cycling using following conditions: a total of 35 PCR cycles each consisting of 30 sec at 94°C, followed by 60 sec at 59°C and 60 sec at 72°C. (4) Prepare 2% (w/v) agarose gel by dissolving 2.0 g agarose (electrophoresis grade) in 100 ml 0.5 × TBE and add 2 µl ethidium bromide per 100 ml gel. (5) Add 10 µl loading dye to each PCR tube and load 15 µl of PCR reaction-dye mixture into each gel well. (6) Analyze PCR products for O157:H7 specific profile by electrophoresis at 100 mV for 30–40 min.

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Escherichia

Notes: (i) Studies have shown that 6 h incubation may suffice in some instances to enable rapid isolation of O157:H7 from foods, but the 18–24 h enrichment was found to be more sensitive and therefore is recommended. (ii) Additional selective enrichment by use of IMS is useful in the analysis of some foods, particularly those with high levels of competing microbial flora such as fecal samples. IMS can be performed using anti-O157 immunomagnetic beads (e.g., Dynabeads) that are commercially available (Dynal Inc., Lake Success, NY), following manufacturer’s instructions. After IMS 150 µl of the resuspended beadbacteria complex can be spread onto a CT-SMAC plate over one quarter of the plate with a sterile swab (this ensures the break-up of the bead-bacteria complexes). Streak for isolation with sterile loops and incubate plates at 37°C for 18–24 h. (iii) Most non-O157:H7 E. coli ferment sorbitol, about 6% of the isolates (atypical strains) do not. These atypical strains along with other sorbitol non-fermenting bacteria such as Morganella and Hafnia appear identical to typical O157:H7 colonies on CT-SMAC agar. Hence confirmatory bacteriological tests must be performed to distinguish these from the O157:H7 isolates. (iv) Alternatively, presumptive colonies can be grown in 2 ml LB broth at 37°C for 6–8 h and gDNA can be isolated from liquid culture using any of the commercial gDNA extraction kits (e.g., DNeasy Tissue Kit, Qiagen, USA).

27.3  Concluding Remarks O157:H7 is capable of causing life threatening illness and represents a serious threat to public health worldwide. The potential for large-scale outbreaks from a wide variety of foods contaminated with O157:H7 and widespread prevalence in animal sources have necessitated the development and evaluation of rapid, sensitive, and specific methods for detection and surveillance for this pathogen. Although conventional microbiological procedures remain an integral part (gold standard) of detection methods, molecular identification of several genetic loci encoding serotype-specific and virulence associated markers has greatly facilitated development of new methods for detection of O157:H7. Use of ELISA assays is also limited by the possible false positive and negative results, although ELISA assays offer a rapid means of screening samples for the presumptive identification of Shiga toxin producing E. coli as well as specific detection of O157 and H7 antigens and thereby reducing the time required for diagnosis, especially in clinical settings. Multiplex PCR and fluorogenic exonuclease assays (e.g., TaqMan PCR) are among the most powerful developments for testing of complex sample matrices. Challenges remain such as the need to

concentrate large sample volumes, and to grapple with low template concentrations in the presence of high CFU of background microflora, and PCR inhibitory substances. The options for detection and confirmation of O157:H7 span both conventional and advanced molecular tools that can be used in a complementary fashion (Figure 27.1). Currently, PCR analysis of primary enrichment cultures appears to be the most sensitive and specific means of screening for the presence of O157:H7. Simultaneous identification of serotype-specific genes as well as multiple virulence factors is, however, necessary for the definitive detection of O157:H7 using PCR. PCR based assays detect chromosomal gene sequences that can be present in viable and dead cells and, therefore, no determination can be made concerning the presence of only viable cells in a sample. This limitation can be partially overcome by the recently developed reverse transcriptase-PCR-based assays or by simultaneous testing of enrichment and nonenrichment samples. Another limitation associated with the use of PCR methods alone for the detection of O157:H7 is that these do not allow recovery of the isolate for epidemiological purposes. Moreover, false positive or false negative reactions could occur if the sample contains mixed microflora that share target genes used for the identification of O157:H7. Coupling PCR with microarrays provides the additional advantage of simultaneously screening for a larger number of target genes with the added benefit of increased confidence because positive detection events are more likely to be real compared with detection of products by agarose gel electrophoresis. Therefore, decisions about the selection of O157:H7 detection methods will involve striking a balance between several factors such as the speed, sensitivity, specificity, availability of proper equipments, skilled personnel and the cost involved in detection procedures. At present, however, comprehensive screening of samples using a combination of pre-enrichment and molecular methods provides a means for rapid and accurate detection of O157:H7, especially in outbreak situations where speed and reliability are critical selection criteria.

Acknowledgments This work was funded in part by the National Institute of Allergy and Infectious Diseases, National Institutes of Health, Department of Health and Human Services, under contract no. N01-AI-30055, and by the Agricultural Animal Health Program and Washington State University.

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387 136. Hu, Y., Zhang, Q. and Meitzler, J.C. Rapid and sensitive detection of Escherichia coli O157:H7 in bovine faeces by a multiplex PCR. J. Appl. Microbiol., 87, 867, 1997. 137. Campbell, G.R. et al. Detection of Escherichia coli O157:H7 in soil and water using multiplex PCR. J. Appl. Microbiol., 91, 1004, 2001. 138. Fratamico, P.M., Bagi, L.K. and Pepe, T. A multiplex polymerase chain reaction assay for rapid detection and identification of Escherichia coli O157:H7 in foods and bovine feces. J. Food Prot., 63, 1032, 2000. 139. Stewart, D.S., Tortorello, M.L. and Gendel, S.M. Evaluation of DNA preparation techniques for detection of the SLT-1 gene of Escherichia coli O157: H7 in bovine faeces using the polymerase chain reaction. Lett. Appl. Microbiol., 26, 93, 1998. 140. McKillip, J.L., Jaykus, L.A. and Drake, M.A. A comparison of methods for the detection of Escherichia coli O157:H7 from artificially-contaminated dairy products using PCR. J. Appl. Microbiol., 89, 49, 2000. 141. Johnston, L.M. et al. A simple method for the direct detection of Salmonella and Escherichia coli O157:H7 from raw alfalfa sprouts and spent irrigation water using PCR. J. Food Prot., 68, 2256, 2005. 142. Tortorello, M.L. et al. Comparison of methods for determining the presence of Escherichia coli O157:H7 in apple juice. J. Food Prot., 61, 1425, 1998. 143. Fitzmaurice, J. et al. Comparison of a membrane surface adhesion recovery method with an IMS method for use in a polymerase chain reaction method to detect Escherichia coli O157:H7 in minced beef. J. Microbiol. Methods, 59, 243, 2004. 144. Islam, M.A. et al. Evaluation of immunomagnetic separation and PCR for the detection of Escherichia coli O157 in animal feces and meats. J. Food Prot., 69, 2865, 2006. 145. Visetsripong, A. et al. Detection of Escherichia coli O157: H7 vt and rfb(O157) by multiplex polymerase chain reaction. Southeast Asian J. Trop. Med. Public Health, 38, 82, 2007. 146. Tyagi, S. and Kramer, F.R. Molecular beacons: probes that fluoresce upon hybridization. Nat. Biotechnol., 14, 303, 1996. 147. Belanger, S.D. et al. Rapid detection of Shiga toxin-producing bacteria in feces by multiplex PCR with molecular beacons on the smart cycler. J. Clin. Microbiol., 40, 1436, 2002. 148. Witham, P.K. et al. A PCR-based assay for the detection of Escherichia coli Shiga-like toxin genes in ground beef. Appl. Environ. Microbiol., 62, 1347, 1996. 149. Reischl, U. et al. Real-time fluorescence PCR assays for detection and characterization of Shiga toxin, intimin, and enterohemolysin genes from Shiga toxin-producing Escherichia coli. J. Clin. Microbiol., 40, 2555, 2002. 150. Oberst, R.D. et al. PCR-based DNA amplification and presumptive detection of Escherichia coli O157:H7 with an internal fluorogenic probe and the 5’ nuclease (TaqMan) assay. Appl. Environ. Microbiol., 64, 3389, 1998. 151. Sharma, V.K., Dean-Nystrom, E.A. and Casey, T.A. Semiautomated fluorogenic PCR assays (TaqMan) forrapid detection of Escherichia coli O157:H7 and other shiga toxigenic E. coli. Mol. Cell. Probes, 13, 291, 1999. 152. Fu, Z., Rogelj, S. and Kieft, T. L. Rapid detection of Escherichia coli O157:H7 by immunomagnetic separation and real-time PCR. Int. J. Food Microbiol., 99, 47, 2005. 153. Artz, R.R. et al. Potential pitfalls in the quantitative molecular detection of Escherichia coli O157:H7 in environmental matrices. Can. J. Microbiol., 52, 482, 2006.

388 154. Holicka, J. et al. A rapid (one day), sensitive real-time polymerase chain reaction assay for detecting Escherichia coli O157:H7 in ground beef. Can. J. Microbiol., 52, 992, 2006. 155. Fortin, N.Y., Mulchandani, A. and Chen, W. Use of real-time polymerase chain reaction and molecular beacons for the detection of Escherichia coli O157:H7. Anal. Biochem., 289, 281, 2001. 156. Ibekwe, A.M. et al. Multiplex fluorogenic real-time PCR for detection and quantification of Escherichia coli O157:H7 in dairy wastewater wetlands. Appl. Environ. Microbiol., 68, 4853, 2002. 157. Sharma, V.K. and Dean-Nystrom, E.A. Detection of enterohemorrhagic Escherichia coli O157:H7 by using a multiplex real-time PCR assay for genes encoding intimin and Shiga toxins. Vet. Microbiol., 93, 247, 2003. 158. Spano, G. et al. Real-time PCR for the detection of Escherichia coli O157:H7 in dairy and cattle wastewater. Lett. Appl. Microbiol., 40, 164, 2005. 159. Abu Al-Soud, W. and Radstrom, P. Effects of amplification facilitators on diagnostic PCR in the presence of blood, feces, and meat. J. Clin. Microbiol., 38, 4463, 2000. 160. Akane, A. et al. Identification of the heme compound copurified with deoxyribonucleic acid (DNA) from bloodstains, a major inhibitor of polymerase chain reaction (PCR) amplification. J. Forensic Sci., 39, 362, 1994. 161. Monteiro, L. et al. Complex polysaccharides as PCR inhibitors in feces: Helicobacter pylori model. J. Clin. Microbiol., 35, 995, 1997. 162. Wilson, I.G. Inhibition and facilitation of nucleic acid amplification. Appl. Environ. Microbiol., 63, 3741, 1997. 163. Tebbe, C.C. and Vahjen, W. Interference of humic acids and DNA extracted directly from soil in detection and transformation of recombinant DNA from bacteria and a yeast. Appl. Environ. Microbiol., 59, 2657, 1993. 164. von Wintzingerode, F., Gobel, U.B. and Stackebrandt, E., Determination of microbial diversity in environmental samples: pitfalls of PCR-based rRNA analysis. FEMS Microbiol. Rev., 21, 213, 1997. 165. Chandler, D.P. et al. Automated immunomagnetic separation and microarray detection of E. coli O157:H7 from poultry carcass rinse. Int. J. Food Microbiol., 70, 143, 2001. 166. Klerks, M.M., Zijlstra, C. and van Bruggen, A.H. Comparison of real-time PCR methods for detection of Salmonella enterica and Escherichia coli O157:H7, and introduction of a general internal amplification control. J. Microbiol. Methods, 59, 337, 2004. 167. McKillip, J.L., Jaykus, L.A. and Drake, M. Nucleic acid persistence in heat-killed Escherichia coli O157:H7 from contaminated skim milk. J. Food Prot., 62, 839, 1999. 168. Yaron, S. and Matthews, K.R. A reverse transcriptase-polymerase chain reaction assay for detection of viable Escherichia coli O157:H7: investigation of specific target genes. J. Appl. Microbiol., 92, 633. 2002. 169. Kushner, S.R. mRNA decay. In Escherichia coli and Salmonella cellular and molecular biology. Neidhardt, F.C., Ed. American Society of Microbiology: Washington DC, 1996, p. 849. 170. McIngvale, S.C., Elhanafi, D. and Drake, M.A. Optimization of reverse transcriptase PCR to detect viable Shiga-toxinproducing Escherichia coli. Appl. Environ. Microbiol., 68, 799, 2002.

Molecular Detection of Foodborne Pathogens 171. Fitzmaurice, J. et al. Application of real-time PCR and RT-PCR assays for the detection and quantitation of VT 1 and VT 2 toxin genes in E. coli O157:H7. Mol. Cell. Probes, 18, 123, 2004. 172. Sharma, V.K. Real-time reverse transcription-multiplex PCR for simultaneous and specific detection of rfbE and eae genes of Escherichia coli O157:H7. Mol. Cell. Probes, 20, 298, 2006. 173. Tsai, T.Y. et al. Detection of viable enterohemorrhagic Escherichia coli O157 using the combination of immunomagnetic separation with the reverse transcription multiplex TaqMan PCR system in food and stool samples. J. Food Prot., 69, 2320, 2006. 174. Liu, Y. et al. Detection of viable but nonculturable Escherichia coli O157:H7 bacteria in drinking water and river water. Appl. Environ. Microbiol., 74, 1502, 2008. 175. Weagant, S., Bryant, J. and Jeinneman, K. An improved rapid technique for isolation of Escherichia coli O157:H7 from foods. J. Food Prot., 58, 7, 1994. 176. Call, D.R. Challenges and opportunities for pathogen detection using DNA microarrays. Crit. Rev. Microbiol., 31, 91, 2005. 177. Call, D.R., Borucki, M.K. and Loge, F.J. Detection of bacterial pathogens in environmental samples using DNA microarrays. J. Microbiol. Methods, 53, 235, 2003. 178. Al-Khaldi, S.F. et al. DNA microarray technology used for studying foodborne pathogens and microbial habitats: minireview. J. AOAC Int., 85, 906, 2002. 179. Kuipers, O.P. et al. DNA-microarrays and food-biotechnology. Antonie Van Leeuwenhoek, 76, 353, 1999. 180. Harrington, C.A., Rosenow, C. and Retief, J. Monitoring gene expression using DNA microarrays. Curr. Opin. Microbiol., 3, 285, 2000. 181. Call, D.R., Brockman, F.J. and Chandler, D.P. Detecting and genotyping Escherichia coli O157:H7 using multiplexed PCR and nucleic acid microarrays. Int. J. Food Microbiol., 67, 71, 2001. 182. Jin, H.Y. et al. Microarray analysis of Escherichia coli O157:H7. World J. Gastroenterol., 11, 5811, 2005. 183. Chizhikov, V. et al. Microarray analysis of microbial virulence factors. Appl. Environ. Microbiol., 67, 3258, 2001. 184. Jin, D.Z. et al. Detection and identification of enterohemorrhagic Escherichia coli O157:H7 and Vibrio cholerae O139 using oligonucleotide microarray. Infect. Agent Cancer, 2, 23, 2007. 185. Kim, H. et al. A molecular beacon DNA microarray system for rapid detection of E. coli O157:H7 that eliminates the risk of a false negative signal. Biosens. Bioelectron., 22, 1041, 2007. 186. Gonzalez, S.F. et al. Simultaneous detection of marine fish pathogens by using multiplex PCR and a DNA microarray. J. Clin. Microbiol., 42, 1414, 2004. 187. Warsen, A.E. et al. Simultaneous discrimination between 15 fish pathogens by using 16S ribosomal DNA PCR and DNA microarrays. Appl. Environ. Microbiol., 70, 4216, 2004. 188. Lane, S. et al. Amplicon secondary structure prevents target hybridization to oligonucleotide microarrays. Biosens. Bioelectron., 20, 728, 2004. 189. Feng, P. and Weagant, S.D. Diarreagenic E. coli. Bacteriological analytical manual available online at http://www.cfsan.fda. gov/~ebam/bam-4a.html 2002.

Escherichia 190. Arthur, T.M. et al. Comparison of the molecular genotypes of Escherichia coli O157:H7 from the hides of beef cattle in different regions of North America. J. Food Prot., 70, 1622. 2007. 191. LeJeune, J.T. and Christie, N.P. Microbiological quality of ground beef from conventionally-reared cattle and “raised without antibiotics” label claims. J. Food Prot., 67, 1433, 2004. 192. Awais, R. et al. Occurrence of virulence genes associated with enterohemorrhagic Escherichia coli in raw municipal sewage. Biochem. Eng. J., 33, 53, 2007.

389 193. Bellin, T. et al. Rapid detection of enterohemorrhagic Escherichia coli by real-time PCR with fluorescent hybridization probes. J. Clin. Microbiol., 39, 370, 2001. 194. Sharma, V.K. Detection and quantitation of enterohemorrhagic Escherichia coli O157, O111, and O26 in beef and bovine feces by real-time polymerase chain reaction. J. Food Prot., 65, 1371, 2002. 195. Hsu, C.F., Tsai, T.Y. and Pan, T.M. Use of the duplex TaqMan PCR system for detection of Shiga-like toxin-producing Escherichia coli O157. J. Clin. Microbiol., 43, 2668, 2005.

28 Klebsiella

Beatriz Meurer Moreira, Marco Antonio Lemos Miguel, and Angela Christina Dias de Castro Federal University of Rio de Janeiro

Maria Silvana Alves

Federal University of Juiz de Fora

Rubens Clayton da Silva Dias University of California

Contents 28.1 Introduction.................................................................................................................................................................... 391 28.1.1 Taxonomy of the Genus Klebsiella................................................................................................................... 391 28.1.2 Natural Habitat and Epidemiology of Klebsiella.............................................................................................. 392 28.1.3 Community-Acquired Klebsiella Infections..................................................................................................... 392 28.1.4 Healthcare Associated Infections caused by Klebsiella................................................................................... 393 28.1.5 Klebsiella as an Enteropathogen....................................................................................................................... 394 28.1.6 Klebsiella as a Foodborne Pathogen................................................................................................................. 394 28.1.7 Isolation and Biochemical Identification.......................................................................................................... 394 28.1.8 Molecular Identification.................................................................................................................................... 394 28.1.9 Molecular Typing.............................................................................................................................................. 395 28.2 Methods.......................................................................................................................................................................... 396 28.2.1 Reagents and Equipment................................................................................................................................... 397 28.2.2 Sample Collection and Preparation.................................................................................................................. 397 28.2.2.1 Sample Processing and Culture ...................................................................................................... 397 28.2.2.2 DNA Isolation.................................................................................................................................. 397 28.2.3 Detection Procedures........................................................................................................................................ 398 28.2.3.1 PCR Amplification........................................................................................................................... 398 28.2.3.2 Sequence Analysis of the rpoB Gene Fragments............................................................................ 399 28.3 Conclusions and Future Perspectives............................................................................................................................. 399 Acknowledgments...................................................................................................................................................................... 401 References.................................................................................................................................................................................. 401

28.1  Introduction 28.1.1 Taxonomy of the Genus Klebsiella The designation of the genus Klebsiella was a tribute to the German microbiologist Edwin Klebs in 1885. Historically, the medical importance of this genus has led to the recognition of three species: Klebsiella pneumoniae, responsible for pneumonia; Klebsiella ozaenae, the cause of ozena (chronic atrophic rhinitis); and Klebsiella rhinoscleromatis, the cause of rhinoscleroma (chronic granulomatous disease), as described in extensive reviews.1,2 In the latest edition (1984) of Bergey's Manual of Systematic Bacteriology, based on phenotypic characteristics and data derived from DNA–DNA hybridization studies,

the genus Klebsiella was classified into five species: K. pneumoniae, Klebsiella oxytoca, Klebsiella terrigena, Klebsiella ornithinolytica, and Klebsiella planticola. K. pneumoniae comprised three subspecies: K. pneumoniae subsp. pneumoniae, K. pneumoniae subsp. ozaenae, and K. pneumoniae subsp. rhinoscleromatis.3 However, with increasing use of DNA–DNA hybridization techniques and additional molecular studies, new species have been described and others reclassified or moved to the new genus Raoultella. In the current text, the term Klebsiella refers to Klebsiella and Raoultella species, unless specified differently. Thus, by sequencing of 16S rRNA and phoE genes, Calymmatobacterium granulomatis, the supposed agent of donovanosis, was renamed Klebsiella granulomatis; based 391

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on 16S rRNA and rpoB sequence analysis, the new genus Raoultella was proposed to accommodate the three species previously called K. planticola, K. terrigena, and K. ornithinolytica while K. oxytoca was shown to form a distinct genogroup; two new Klebsiella species were described: Klebsiella variicola, from analysis of rpoB, gyrA, mdh, infB, phoE, and nifH sequences and Klebsiella singaporensis, based on 16S rRNA genes and rpoB gene sequences; and finally, 16S rRNA gene sequences for two proposed species, Klebsiella milletis and Klebsiella senegalensis, and 16S rRNA and rpoB sequences for the proposed species Klebsiella alba were deposited in the GenBank (http://www.ncbi.nlm.nih.gov) in 2003 and 2007, respectively.4–7 However, no data has been published describing these three last species to date. A summary of the classification of Klebsiella is presented in Table 28.1. Subgroups within species were also determined. Based on nucleotide variations of the gyrA, parC, and rpoB genes, clinical isolates of K. pneumoniae fall into four phylogenetic groups named KpI, KpII-A, KpII-B, and KpIII.8,11,13 The species K. variicola corresponds to group KpIII.6 In addition, K. oxytoca has been shown to include five distinct phylogenetic lines (KoI, KoII, KoIII, KoIV, and KoVI) identified by 16S rRNA, rpoB, gyrA, gapDH, and blaOXY gene sequencing.15

28.1.2 Natural Habitat and Epidemiology of Klebsiella Klebsiella is ubiquitous in nature, and has been found in surface water, sewage, soil, and plants, and colonize mucosal

surfaces of mammals such as humans, horses, and swine.1 The investigation of natural surface water (freshwater, salt water, and brackish water) in Germany led to the isolation of Klebsiella species from 110 samples from streams, lakes, and the Baltic Sea in various geographic areas; K. pneumoniae was detected in 52%, K. oxytoca in 27%, and Raoultella planticola in 22% of the samples.17 K. pneumoniae causes serious infections in humans.18 It is reported also as an animal pathogen, and recognized as a main cause of clinical mastitis in cows, resulting in high milk losses and mortality of the affected animal.19 The presence of virulence traits in K. pneumoniae from water and contemporary human clinical isolates has been shown to be comparable, indicating environmental isolates are potential sources of significant pathogens for humans.17,20 Fecal shedding of K. pneumoniae by cows contributes to the presence of a variety of strains in dairy herds, which may in turn be a source of colonization for humans19.

28.1.3 Community-Acquired Klebsiella Infections K. pneumoniae has been recognized as cause of communityacquired pneumonia (CAP) since its first description, more than 120 years ago. The clinical presentation of K. pneumoniae CAP is dramatic: sudden onset, high fever, hemoptysis, and toxemia. Chest radiograph abnormalities such as bulging interlobar fissure and cavitary abscesses are prominent.21 Early initiation of an adequate antibiotic therapy has been

Table 28.1 Classification of Klebsiella and Raoultella Species

Subspecies

K. pneumoniae

K. pneumoniae subsp pneumoniae

K. granulomatis K. variicola K. singaporensis K. milletis K. senegalensis K. alba

K. pneumoniae subsp. ozaenae K. pneumoniae subsp. rhinoscleromatis * – – – – –

Genomospecies K. oxytoca

–

R. planticola R. terrigena R. ornithinolytica

– – –

Phylogenetic Group KpI KpII-A KpII-B KpIII KpI KpI – KpIII – – – – KoI KoII KoIII KoIV KoVI – – –

Reference 3,5,8–10 3,5,8,9,11 3,5,8–11 3,5,8–10 3,5,8–10 3,5,8–10 4,5,9,12 6,8,10,13 7 AY217656 AY217655 GenBank accession Nos. EU301692 and EF154517 3,5,8–11,14 3,5,8–11,14,15 3,5,15,16 3,5,15 3,515 5 5 5

–: Not determined. *: Some authors have suggested K. granulomatis belongs to one of the K. pneumoniae groups, or to a new cluster.4,8,12

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shown to improve the prognosis; however, CAP still remains one important cause of death, especially in the elder and those with underlying diseases.22 An emerging and invasive presentation of K. pneumoniae infection is primary pyogenic liver abscess (PLA).21 K. pneumoniae PLA differs from traditional liver abscess because it is community-acquired, patients have no history of hepatobiliary diseases, and may present other septic metastatic lesions.21,23 Most reports originate from Taiwan and Southeast Asia, but this disease has now been described also in Europe, North America, and Australia.24–26 Apparently, there is a geographical preponderance of a syndrome consisting of liver abscess, meningitis, and endophtalmitis in Taiwan.26 Although the reasons for the geographical diversity are unknown, genetic susceptibility may be important.27 The disease has been caused by K1 capsular serotype hypermucoviscous (HMVKp) strains. A main virulence factor in HMVKp strains is magA, a gene encoding an iron transport protein (originally described in Magnetospirillum sp.), responsible for the hypermucoviscous phenotype.25 Genotyping of K1 K. pneumoniae isolates from three different continents by pulsed field gel electrophoresis (PFGE) and multilocus sequence typing (MLST) revealed that a major clone causes the disease; isolates belonging to a main PFGE cluster, with one exception, belonged to ST23.26 K. pneumoniae is also an important cause of community-acquired central nervous system infection in East Asia, despite the presence of a predisposing condition. Meningitis and brain abscess were reported from patients in Singapore, some of them with diabetes mellitus.28 A strong association between community-acquired bacterial meningitis due to K. pneumoniae and liver cirrhosis in adult patients was reported.29 Another serious Klebsiella community-acquired infection is rhinoscleroma caused by K. pneumoniae subsp. rhinoscleromatis. This is a chronic granulomatous infectious disease that may present lesions in the respiratory tract. The nasal cavity is most often affected, but lesions may also involve the larynx, oral cavity, paranasal sinuses, soft tissue of the lips and nose, trachea, bronchi, and the orbit or middle ear.30 Endemic regions are tropical Africa, India, Southeast Asia, Central, and South America, and Central Europe.31 The incidence of the disease in the US population may be on the rise with migration and travel abroad. The diagnosis in nonendemic areas is often delayed due to unfamiliarity with the disease, stage-dependent clinical and histological findings, and the fact that about 40% of cultures are negative for K. rhinoscleromatis. Although this disease is rarely lethal, late diagnosis may lead to nasal and airway obstruction and nasal deformity from erosive processes.32 K. pneumoniae subsp. ozaenae is the etiologic agent of ozena, a primary atrophic rhinitis. Ozena is characterized by mucosal atrophy together with bone resorption and a fetid thick endonasal crust.33 Osseous wall thickening of the maxillary and ethmoid sinuses has also been evidenced.34

28.1.4 Healthcare Associated Infections caused by Klebsiella Between 1997 and 2002, Klebsiella was responsible for 7–10% of all healthcare-associated bloodstream infections in Europe, Latin America, and North America, as reported by the SENTRY Antimicrobial Surveillance Program.35 Immunocompromised patients, especially the elderly and infants, are the population at most risk for infections.36 K. pneumoniae may cause a variety of healthcare-associated infections, including bloodstream-infections, ventilatorassociated pneumonia, meningitis, soft tissue, and urinarytract infections.1 Endemic infections and outbreaks have plagued hospitalized patients, usually in intensive care-units, where the neonate is especially susceptible.37 K. oxytoca is the second most frequent Klebsiella isolate recovered in hospitals. R. planticola is seen as an aquatic, botanic, and soil bacterium, but has been rarely isolated from clinical specimens, and may cause bloodstream infection and pancreatitis.38 Colonization of the gastrointestinal tract of patients usually precedes infection.39–45 In fact, the gastrointestinal tract constitutes a main reservoir for the spread of these organisms on hands and medical equipment to uncolonized patients admitted to hospitals. Klebsiella isolates have been known to be well adapted to the hospital environment. These organisms can survive longer than other enteric bacteria on hands and hospital environmental surfaces, facilitating cross-infection within hospitals.37,46,47 Poor hand hygiene by healthcare personnel is probably a main issue in the spread of these nosocomial pathogens between patients. Other environmental sources are important; for example, contamination of enteral feeding by K. oxytoca has been shown to be the source of throat and stool colonization of premature babies.43 The strategy to decrease the prevalence of healthcare-associated infections caused by Klebsiella isolates should include restriction of antimicrobial use, particularly third-generation cephalosporins, and the reinforcement of the correct employment of barrier precautions.40,43,46 The effectiveness of infection control measures is enhanced by the early detection of patients whose gastrointestinal tracts have been colonized.45,48 Antimicrobial resistance represents a serious problem in this bacterial group. The prevalence of isolates that produce extended-spectrum β-lactamase (ESBL), which hydrolyses broad-spectrum cephalosporins, monobactams, and penicillins, is increasing worldwide due to the spread of mobile genetic elements and the dispersion of epidemic clones.49 ESBL-producing strains are often resistant to aminoglycosides, sulfonamides, and fluoroquinolones.50 The escape of K. pneumoniae strains from hospitals to the community poses a serious challenge to the treatment of communityacquired infections.23 The frequency of ESBL-production by community-acquired K. pneumoniae isolates varies in different countries; however, these strains already emerged in Europe, Asia, and South and North America.23,51–53 Carbapenem-resistant K. pneumoniae is an additional serious problem observed in hospitals, but there is little

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doubt that these other resistant strains will appear in the community.23 With the rise in the ­number resistant infections, the mortality of K. pneumoniae has increased and treatment has become more complicated.37,54

28.1.5  Klebsiella as an Enteropathogen Klebsiella is an accidental cause of diarrhea. A study of a gastroenteritis outbreak revealed that an LT-enterotoxin producing K. pneumoniae capsular serotype K15 caused the diarrhea, and the outbreak source was contaminated turkey, kept at improper temperatures for a long time.55 In fact, the isolation of LT- and ST-enterotoxin producing K. pneumoniae has been described since 1975.56,57 Another enteropathogenic mechanism of K. pneumoniae is enteroinvasive activity: a case of severe sepsis followed the ingestion a hamburger heavily contaminated by an LT-producing isolate.58 Although K. pneumoniae has not been fully characterized as an enteropathogen, by analogy with diarrheagenic E. coli, three distinct patterns of adhesion to epithelial cells have been described: diffuse, aggregative, and localized.59,60 The pathogenic significance of these adhesion patterns remains to be determined. K. oxytoca may be also an enteropathogen, identified as an emerging cause of antibiotic-associated hemorrhagic colitis (AAHC).61 Strong evidence of this association has been provided, where hemorrhagic colitis followed administration of amoxicillin with or without clavulanate, metronidazole or clarithromycin to five young healthy outpatients. Spontaneous resolution of symptoms was observed in all cases 3–7 days after stopping the antimicrobial therapy. K. oxytoca stool carriage may be an important factor in the development of AAHC.

28.1.6  Klebsiella as a Foodborne Pathogen Klebsiella is widespread in the environment and the presence of this organism in water or food does not necessarily indicate a pathogenic potential. However, the finding of Klebsiella in food may be of significance, for several reasons. First, Klebsiella is a well known cause of gastroenteritis, although only when heavy contamination occurs. On the other hand, the presence of Klebsiella may severely deteriorate the food.55,58 For example, production of thermal-stable protease has been described in K. oxytoca isolated from raw milk.62 Heat-stable protease may constitute a serious problem to ultra-high temperature (UHT) processed milk, leading to undesirable physical and sensory alterations. Secondly, K. oxytoca may cause antibiotic-associated colitis that can be acquired from various sources, including food.61 Finally, the presence of Klebsiella in food may be a source of gastrointestinal colonization for patients admitted to hospitals, which is, in turn, the first step for the development of systemic infection. Klebsiella may be detected in food or water as a “coliform,” together with other main enteropathogens, such as some strains of E. coli. There are no systems to detect specifically

Molecular Detection of Foodborne Pathogens

Klebsiella in food or water. Detection of ­“coliforms” has been used as an indicator of water pollution and food contamination; however, the finding of Klebsiella does not necessarily indicate fecal contamination. If Klebsiella is suspect of a foodborne disease, determination of bacterial counts in the suspect food source, genotyping of organisms isolated from food and patient’s gastrointestinal specimens, and a full epidemiological investigation are necessary to imply any causal relationship.

28.1.7 Isolation and Biochemical Identification Klebsiella is frequently isolated in the clinical laboratory, where these organisms can be obtained from blood, urine, respiratory secretions and other specimens. Standard laboratory media are suitable for the isolation of this organism.18 MacConkey agar is an excellent media, because it allows for the clear observation of the typical appearance of mucoid or rough colonies. Biochemical identification must include several tests, usually with the help of media that combine some of them. The correct identification of Klebsiella species is not easily accomplished in most clinical microbiology laboratories, because several species share a similar biochemical profile. A suggested scheme propose the following profile for isolates belonging to the most common species of the genera Klebsiella and Raoultella: nonmotile Gram-negative bacilli; glucose fermenters; form typical red colonies on MacConkey agar; show negative oxidase, H2S, arginine, ornithine, and phenylalanine reactions; and citrate use.10 Isolates which are negative for indole production, unable to assimilate histamine and d-melezitose or to grow at 10°C are identified as K. pneumoniae or K. variicola. Possible K. oxytoca isolates are positive for indole production, lysine decarboxylation, Voges-Proskauer test, and growth at 10°C and negative for histamine assimilation. R. planticola are positive for indole production, histamine assimilation, and growth at 10°C and negative for ornithine and d-melezitose. However, currently, a complete identification of all Klebsiella species is possible only with molecular tests. For example, K. pneumoniae and K. variicola can not be confidently separated by biochemical methods, and K. granulomatis is not cultivable in artificial laboratory media.4,10 In addition, some of the tests necessary for identification of species or are not available or are not standardized in most laboratories.

28.1.8 Molecular Identification The study of bacterial taxonomy and phylogeny evolved robustly by comparison of 16S rRNA gene or spacer sequences, and several protocols for molecular identification of pathogens were developed based on analysis of these genes. One of the few studies addressing the molecular detection of foodborne Klebsiella described a PCR method based on 16S–23S rDNA internal transcribed spacer of K. pneumoniae.63 Speciesspecific internal transcribed spacer regions are multicopy non-coding sequences interspaced among conserved bacterial

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and fungal rDNA. The technique can be performed by using one of two pairs of K. pneumoniae specific primers (Pf/Pr1 or Pf/Pr2) (Pf [5′-ATTTGAAGAGGTTGCAAACGAT-3′], Pr1 [5′-TTCACTCTGAAGTTTTCTTGTGTTC-3′], and Pr2 [5′-CCGAAGATGTTTCACTTCTGATT-3′]). The PCR sensitivity was > 102 bacterial colony forming units (CFU)/100 g of infant formula with primers Pf/Pr2, and > 103 bacterial CFU/100 g of infant formula with primers Pf/Pr1. This sensitivity was assessed in serial dilutions of DNA extracted from a bacterial suspension. In addition, the sensitivity of the assay was assessed after pre-enrichment of artificially contaminated formula during two steps of overnight incubation: one in buffered peptone water and another in EE broth (buffered glucose, brilliant green, bile broth). After this additional procedure, the PCR assay was able to detect 1.5 CFU of K. pneumoniae/100 g of infant formula. Since K. pneumoniae and other pathogens known to cause illness in infants have been found in low levels in powdered infant formula,64 preenrichment is probably an important step for this method. The Food and Agriculture Organization of the United Nations has recommended that bacterial counts of coliform organisms in good-quality powdered infant formulas should be of <3 CFU/g. However, milk powders free of Enterobacteriaceae species might offer extra protection to the newborn and especially to the premature baby, because some bacterial multiplication during preparation and storage of contaminated milk does occur. In addition, hygienic precautions are also essential measures to control milk contamination. The analysis of 16S rRNA sequences may have limitations: the presence of several and variable copies of ribosomal coding genes have been described in K. variicola.6 Thus, sequence analysis of several additional housekeeping genes have been used for identification of Klebsiella, including rpoB (RNA polymerase β-subunit), gyrA (DNA gyrase A subunit), mdh (malate dehydrogenase), infB (translation initiation factor 2), phoE (phosphoporine E), and nifH (nitrogenase reductase).4–7 The comparison of the first rpoB partial sequences of Enterobacteriaceae species revealed that divergence levels of sequences were clearly superior to those of 16S rRNA genes.5,65 Additionally, phylogenetic trees of Enterobacteriaceae isolates obtained with rpoB fit better with the classification of these organisms than 16S rRNA trees. Therefore, rpoB sequencing has been frequently used as a means of identification and phylogenetic analysis of enterobacteria. On the other hand, the study of the rpoB gene has not been straightforward. For example, five different primers were used when analysis of this gene was proposed for bacterial identification: two for rpoB amplification and three for sequencing.65 In addition, utilization of the proposed CM7 and CM31b primer pair for rpoB amplification can result in more than one band making the purification of PCR product for sequencing more difficult.10,65 In the present chapter, the authors have chosen to describe in detail rpoB sequencing as a simple molecular approach for the identification of Klebsiella, and describe new primers that facilitate this analysis. However, the initial steps of bacterial isolation and

biochemical identification are still necessary, at least at the Enterobacteriaceae level. New promising approaches have been recently described for detection of Klebsiella in blood culture samples.66–68 One technique involves fluorescence in situ hybridization (FISH) with peptide nucleic acid (PNA) probes targeting rRNA genes. The PNA probe targets the K. pneumoniae 23S rRNA gene, and preliminary data suggest it provides an accurate diagnosis within 3 h. Another sensible and specific technique involves real-time PCR. In one study, a system by Roche Molecular Biochemicals (LightCycler) using a 16S rRNA gene K. pneumoniae specific probe was 100% sensitive and specific for positive blood cultures, which included Pseudomonas aeruginosa, E. coli and K. oxytoca as negative controls.67 A quantitative estimate is possible with this assay, which was able to detect as few as ten bacterial cells. The assay was complete in 2 h. A second study on real-time PCR assay (LightCycler SeptiFast Test M Grade, Roche Molecular Systems) described the test performed directly from blood samples.68 The assay targets specific rDNA internal transcribed spacer regions. The test was complete in 6 h, and detects 20 pathogens (or groups of pathogens), but K. pneumoniae is not differentiated from K. oxytoca. In the study described,68 the PCR test detected more pathogens than the conventional blood culture system; however, no samples were positive for Klebsiella (pneumoniae/oxytoca) in the patient population studied. Whether clinical laboratories will be able to cope with the costs of routine molecular approaches is yet to be determined. In addition, in the clinical laboratory, regular cultures were still necessary to perform the susceptibility tests.

28.1.9 Molecular Typing Molecular typing of pathogens has become an important tool for the epidemiologic investigation of infectious diseases. Different molecular typing methods have enhanced our understanding of the epidemiology of several bacterial infectious diseases, by helping to determine modes of transmission of pathogens, the source and the risk factors for infections. Strain typing methods are used to address two different kinds of problems: to determine if isolates recovered from a localized outbreak of disease belong to a single or to different strains (short term or local epidemiology), and to determine if strains causing disease in one geographic area are related to others obtained world-wide (long term or global epidemiology). Molecular strain typing methods together with classic epidemiology studies have been essential for the understanding of the transmission dynamics of infections caused by Klebsiella.69,70 For instance, a molecular epidemiology study led to the identification of a contaminated ultrasonography coupling gel container of an emergency room as the source of a hospital outbreak.71 Strain typing by PFGE revealed that patient isolates and an isolate form the ultrasonography gel belonged to a single strain. This investigation disclosed two modes of K. pneumoniae transmission: ultrasound scanning by transvaginal ultrasonography, and

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peripartum colonization of neonates, and led to the complete control of the outbreak. In the food industry, bacterial strain typing is being increasingly used to identify sources of infection, product contamination, and to elucidate routes of transmission of pathogens or spoilage organisms. However, information about the molecular epidemiology of Klebsiella is still limited. For example, despite the increasing incidence of clinical mastitis by K. pneumoniae in dairy herds and the impact of infections in cow health and productivity, few studies using molecular techniques have characterized this pathogen. The study of increasing numbers of severe cases of clinical mastitis in a NewYork State dairy herd in 2006 disclosed interesting data.19 K. pneumoniae was obtained in ten of 17 positive cultures from 20 cows. Bedding samples and animal feces were also analyzed in the search of a potential source. Strain typing by random amplified polymorphic DNA (RAPD) analysis revealed that clinical mastitis was caused by a predominant strain type, which indicated that contagious transmission of K. pneumoniae from cow to cow occurred. Most probably, the RAPD epidemic type caused infection in the index case and subsequently disseminated to other cows via the milking machine or bedding material. Several measures to contain spread of the epidemic strain led to the control of the outbreak. This outbreak was followed by a second increase in the number of clinical mastitis caused by several different Klebsiella strains, but this time due to exposure to isolates commonly present in the dairy farm environment. Analysis of chromosomal DNA restriction patterns by PFGE is an appropriate and discriminatory technique for typing different microorganisms.69 However, PCR-based techniques such as RAPD, enterobacterial repetitive intergenic consensus-PCR (ERIC-PCR), and repetitive extragenic palindromic-PCR (REP-PCR) analysis are faster and easier to perform,72–74 and provide useful discriminatory powers. For typing K. pneumoniae, up to two band differences is a functional cutoff, with ERIC-PCR being the most reliable of the methods.74 PFGE and PCR-based methods have the disadvantage of relying on comparison of banding patterns generated by gel electrophoresis, which is difficult to standardize. Therefore, comparability of patterns within and between laboratories requires strict adherence to established protocols. Even though, reproducibility is not always enough and normalization of patterns is complex. To overcome the poor portability problems of electrophoresis banding techniques, an appealing, sequence-based alternative was developed: MLST. Unfortunately, the applicability of MLST to different pathogens in specific epidemiologic contexts is not yet well understood. MLST is based on the nucleotide sequence of internal regions of housekeeping loci; multiple loci with only minimal nucleotide changes due to conserved protein function are explored. Small sequence variations allow for the assignment of alleles, and the combination of alleles for several loci provides an allele profile that determined a sequence type (ST).

Molecular Detection of Foodborne Pathogens

Subsequently, STs are grouped into clonal complexes based on similarity to a central allelic profile. MLST has been proved useful for local and global epidemiologic studies. In 2005, Diancourt and colleagues75 developed an MLST scheme for K. pneumoniae, available at the http://pubmlst.org/kpneumoniae website. The scheme uses internal fragments of seven housekeeping genes: rpoB, gapA (glyceraldehyde 3-phosphate dehydrogenase), mdh, pgi (phosphoglucose isomerase), phoE, infB, and tonB (periplasmic energy transducer). The analysis of nosocomial isolates with this protocol revealed a high (96%) discriminatory power, compared to ribotyping. Interesting data about Klebsiella infections have been provided by studies using MLST. Paulin-Curlee and colleagues76 evaluated the genetic diversity of K. pneumoniae isolated from clinical mastitis cases, to define genotypes most commonly associated with the disease. Authors used the same MLST protocol proposed by Diancourt and colleagues,75 in addition to PFGE and REP-PCR. A significant genetic diversity among the isolates was observed regardless of the fingerprinting method used: Simpson’s diversity index was 97.0, 96.1, and 93.5%, when strains were analyzed by MLST, PFGE, and PCR-base DNA fingerprinting, respectively. Noteworthy was that three isolates from mastitis in cows belonged to the same MLST types previously observed among human clinical isolates. This finding indicates that K. pneumoniae isolates of bovine origin may be involved in human disease, and illustrates how the increasing use of portable typing data will impact our knowledge about the molecular epidemiology of infections possibly related to food. In 2008, another MLST scheme for K. pneumoniae was developed to characterize human ESBL-producing K. pneumoniae isolates from an outbreak in a Korean hospital.77 This new MLST scheme uses only five housekeeping genes: rpoB, gyrA, gapA, groEL (GroEL protein), and gyrB (DNA gyrase B subunit), and results were comparable to those obtained by PFGE. Unfortunately, the performance of the five-gene protocol was not compared to that of the seven-gene protocol. It is probable still advisable to use the first protocol because it is described in detail in the MLST website, STs may be freely compared to those in the database, and new STs may be easily deposited. Finally, although the MLST method offers a high discriminatory power and inherent objectivity, this technique is still impractical for large-scale epidemiological studies, and simpler screening sequence-based tests are still needed.

28.2  Methods We present below PCR and sequencing methods based on the rpoB gene to classify Klebsiella isolates. Specifically, enterobacterial rpoB sequences were aligned with the ClustalW Multiple Alignment of BioEdit Sequence Alignment Editor, version 7.0.5.3,78 and primers rubF (5′-CTCTGGGCGATCTGGATA-3′) and rubR (5′-CATGT TCGCACCCATCAA-3′) were designed, corresponding to the rpoB gene regions located at bp 1439–1456 and bp

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650 bp

1 Kb Plus DNA ladder

Negative control

R. planticola ATCC 33531

K. pneumoniae ATCC 700603

E. aerogenes ATCC 13048

E. coli ATCC 11229

E. coli ATCC 25922

1 Kb plus DNA Ladder

650 bp

Figure 28.1  Gel electrophoresis of PCR DNA fragments showing the ~617 bp rpoB gene PCR amplicon. PCR products were separated in a 1.5% agarose gel. Lanes are labeled with the bacterial species used as DNA template. The size of the marker in base pairs is shown in both sides.

2038–2055, respectively, of E. coli K-12 strain MG1655. The expected PCR product is ~617 bp. DNA templates of E. coli, Enterobacter aerogenes, Enterobacter cloacae, K. pneumoniae, and R. planticola isolates from reference collections, clinical, and food isolates have been successfully sequenced and identified by the authors with this primer pair (Figure 28.1).

28.2.1 Reagents and Equipment The required reagents and equipment are shown in Table 28.2

28.2.2 Sample Collection and Preparation Even though genotypic assays give tremendous advantages of speed and simplicity, phenotypic methods for the identification and characterization of foodborne bacteria continue to be widely used. First, culture is frequently the only alternative for the detection of an infrequent foodborne pathogen such as Klebsiella. Secondly, bacterial cells must be viable to cause infection; molecular methods usually work for nonviable cells. Finally, bacterial growth in pure culture is usually necessary for strain typing analysis, which is an essential step of the study of outbreaks or to establish potential relatedness between patient- and food-isolates. 28.2.2.1 Sample Processing and Culture For the microbiological investigation of food contamination by Klebsiella, food or water samples must be representative and properly collected with sterile technique. The appropriate sampling procedure for solid, semi-solid, viscous or liquid food must be determined by using Investigations

Operational Manual.79 Samples must be promptly submitted to the laboratory in the original unopened containers and with the original storage conditions maintained as nearly as possible. If the food is frozen it must be refrigerated (~5ºC) for 18 h before analysis. Several methods may be used to detect and estimate counts of microorganisms in food. The widely used standard plate counts (SPC) is well suitable for the detection of Klebsiella, because colonial morphology can be observed for presumptive bacterial identification. Samples (25 g) are placed in 225 ml of a 1% buffered peptone water solution (2% of sodium citrate should be added for the analysis of cheese) and homogenized with a blender, a Stomacher, which homogenizes specimens in a special plastic bag by the vigorous pounding of two paddles, or a Pulsifier, which creates turbulence on the food samples resulting in the release of microorganisms. Buffered peptone water is used as a nonselective pre-enrichment media in which phosphates buffer the medium, peptone is a source of carbon, nitrogen, vitamins, minerals, and sodium chloride maintains the osmotic balance. Therefore, the neutral pH maintained over the preenrichment period results in repair of injured cells that may be sensitive to low pH. After overnight incubation, 1 ml inoculums of each of the serial dilutions (10 –1, 10 –2, and 10 –3) of the homogenate in 1% buffered peptone water are inoculated in an appropriate selective medium. When choosing the selective culture medium, one should consider whether the intrinsic and extrinsic characteristics of the food sample are stressing for the microorganism or not; the differential-selective media vary greatly in their abilities to support resuscitation and growth of stressed cells of microorganisms. The violet red bile agar (VRBA) has been used as the sole agar medium or as an overlay onto trypticase soy agar for detection and enumeration of coliforms. The first choice is used for organisms not expected to be stressed or damaged, while the second is recommended for recovery of injured cells.80–82 Different presentations (broth vs. agar) of a medium also vary in the ability to recover an organism. Authors have observed that MacConkey broth works better than MacConkey agar for the isolation of microorganisms from environmental swab specimens. Suspect colonies are counted after incubation at 37oC for 24–48 h. Biochemical bacterial identification must follow, as described in Section 28.1.7.10 A liquid specimen may be also analyzed by membrane filtration, a preferred technique when small counts of bacteria are expected. Membranes with a 0.45 μm pore size that retain bacteria, but allow water or diluents to pass, are used. A given sample volume is filtered and the membrane is placed onto an agar plate with appropriate media such as blood agar, or MacConkey agar. The culture is incubated at 37oC for 24–48 h, and bacterial colonies are counted and identified. 28.2.2.2 DNA Isolation The protocol described for DNA isolation and amplification has been tested and used by the authors. Other similar

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Molecular Detection of Foodborne Pathogens

Table 28.2 List of Necessary Reagents, Supplies and Equipment Reagents and Supplies Media – Luria Bertani agar plates or other nonselective media plates – MacConkey agar plates – Luria Bertani broth or other nonselective broth Solutions, buffers, and reagents – Distilled water – DNase/RNase-free sterile distilled water – 10× tris-borate EDTA buffer, pH 8.4 (1 M EDTA) – 10× Taq Buffer (with 15 mM MgCl2) – dNTP set 100 mM (each) – Oligonucleotide primers – Taq DNA polymerase (preferably automated hot start PCR) – Electrophoresis loading dye solution, 6× – Ethidium bromide solution – 1 kb plus DNA ladder – Low mass DNA ladder – Ultrapure agarose – 1% buffered peptone water solution – 1% buffered peptone water solution with 2% sodium Citrate

a b c

Equipment – Refrigerator and freezer – Microwave oven – Bench top centrifuge (preferably refrigerated) – Horizontal laminar flow cabinet or bunsen burner for aseptic practices – Standard lab incubator (30–75ºC) – Autoclave – Stirrer/ heater plate – Laboratory scale – Thermocycler – Horizontal electrophoresis system – Image documentation system with UV transilluminator – Adjustable-volume micropipettor: 0.2–2 µl (P2), 1–10 µl (P10), 2–20 µl (P20), 20–100 µl (P100), 50–200 µl (P200), 200–1000 µl (P1000) – Pipette filler or Pipette pump – Sterile disposable polystyrene pipettes or Reusable autoclavable glass pipettes: 1 ml, 10 ml, and 25 ml – Sterile aerosol-barrier pipette tips (for automatic micropipettor) 10 µl, 200 µl, and 1000 µl sizes – Sterile disposable plastic loops and needles or Nichrome or platinum wire loops/needlesa – Polypropylene microcentrifuge tubes: 0.5 ml and 1.5 ml sizes – 0.2 ml thin-wall PCR tubes – 5 ml sterile disposable cell culture roundbottom polypropylene tubes – Autoclavable bacteriological culture glass tubes (Screw-cap, 150 ml capacity) – PCR purification kit – Autoclavable filtration apparatusb – Nylon filter membranes pore size 0.45 μmb – Stomacher or Pulsifier – Stomacher disposable sterile plastic bagc – Colony counter – Disposable gloves

If nichrome or platinum wire loops/needles are used, Bunsen burner is necessary to sterilize the loop by flaming. Optional equipment. For analysis of solid and semi-solid food samples.

protocols can work. Bacterial isolates are incubated at 36 ± 1ºC on Luria Bertani (LB) agar (or other nonselective media) plates overnight and then a single colony of each isolate is inoculated into 3 ml LB broth (or other nonselective media) and further incubated for 16–18 h at 36 ± 1ºC. A 0.5-ml bacterial suspension is centrifuged at 30,000 × g for 5 min and the supernatant discarded. The bacterial pellet is washed in 200 µl of sterile double distilled water, and the pellet resuspended in 100 µl of sterile double distilled water and boiled for 10 min. The supernatant is recovered after centrifuging at 30,000 × g for 5 min and kept on ice. All steps are done at 4ºC. Bacterial DNA can be kept at –20ºC until use.

28.2.3  Detection Procedures 28.2.3.1  PCR Amplification (1) Allocate 24 µl of PCR mixture (containing 2.5 µl of 10× PCR Buffer, 1.5 mM of MgCl2, 200 µmol each of dATP, dCTP, dGTP, and dTTP, 0.2 µmol of each primer (forward rubF and reverse rubR primers), and 1.25 U of Taq DNA polymerase) into each 0.2 ml PCR tube. (2) Add 1 µl of template DNA into each PCR tube. (3) Conduct PCR amplification in a PCR machine using the following cycling programs: an initial denaturation step at 95°C for 2 min; 33 cycles of

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Klebsiella

denaturation at 95°C for 1 min, primer annealing at 52°C for 1 min, primer extension at 72°C for 2 min; and a final extension step at 72°C for 5 min. (4) After completion of PCR amplification, load 10 µl of PCR products (with 2 µl of 6× loading dye) on a 1.5% agarose gels, electrophorese, stain with ethidium bromide, and photograph under ultraviolet light to confirm the amplification of a specific 617 bp product (Figure 28.1). 28.2.3.2 Sequence Analysis of the rpoB Gene Fragments (1) Purify the 617 bp PCR amplicon using a purification kit. (2) Perform sequencing reactions for both DNA strands by using forward rubF and reverse rubR primers. (3) Align the overlapping sequences of the 512 bp fragments (located at bp 1498–2009 of sequence accession #NC000913 in GenBank database) to create the consensus sequence for analysis. (4) Conduct similarity searches with other sequences in GenBank with the program BLASTn (http://www. ncbi.nlm.nih.gov). (Note: the classification of the isolate can be accomplished by a similarity equal to or greater than 97%.83 The BLASTn similarity search for the 512 bp fragment of strains included in the design of primer rubF and rubR revealed rpoB sequence similarities greater than 98%). (5) After performing the similarity search, a thorough analysis includes the construction of a phylogenetic tree with sequences of key strains of several species, and the confirmation of the clusters of the isolates under investigation. Comments. Figure 28.2 presents an example of tree constructed with rpoB 512 bp fragments corresponding to primers rubF and rubR of 50 strains obtained from the GenBank database. The strains are listed at the end of this section. The multiple alignment program used in this example is the ClustalW Multiple Alignment of MEGA4 software.84 The alignment data were assessed by bootstrap analyses based on 1,000 resamplings. Evolutionary distances were computed by the neighbor-joining distance method using the Kimura 2-parameter method and are in the units of the number of base substitutions per site.85,86 Codon positions included were 1st+2nd+3rd+Non-coding. The tree was drawn to scale, with branch lengths in the same units as those of the evolutionary distances used to infer the phylogenetic tree. There were a total of 512 positions in the final dataset. The significance of the branching order was evaluated by bootstrap analysis with 500 replicates. The phylogenetic tree shows that the 512 bp rpoB gene fragment analysis is able to discriminate the sequences from 20 different species. The strains included in the tree are Citrobacter freundii ATCC11102 (accession #U77434); Enterobacter aerogenes LMG2094 (DQ836222); Enterobacter cloacae ATCC13047T

(AJ543726); E. coli K-12 MG1655 (NC000913), O157:H7 EDL933 (NC002655), ATCC25290, CFT073, and Sakai (U77436, NC004431, and NC002695, respectively); Escherichia fergusonii ATCC35469 (U77437); Hafnia alvei ATCC13337 (U77438); K. oxytoca ATCC13182, K3, K43, K45, K50, K56, K89, K124, and K140 (U77442, DQ415458, DQ415461, DQ415462, DQ415464, DQ415465, DQ415468, DQ415469, and DQ415470, respectively); K. pneumoniae K14 and K18 (DQ415473 and DQ415474, respectively); K. pneumoniae subsp. ozaenae ATCC11296 (AF129445); K. pneumoniae subsp. pneumoniae ATCC13883, ATCC700603, and MGH78578 (U77444, EU693237, and NC009648, respectively); K. pneumoniae subsp. rhinoscleromatis ATCC13884 (AF129446); K. variicola ATCCBAA-830, K27, K40, and K166 (AY367356, DQ415499, DQ415500, and DQ415503, respectively); Kluyvera ascorbata ATCC33433T (U77439); Proteus mirabilis ATCC29906 (U77445); Providencia stuartii ATCC29914 (U77446); R. ornithinolytica ATCC31898 (AF129447); R. planticola ATCC33531, K111, and K112 (AF129449, DQ415504, and DQ415505, respectively); R. terrigena ATCC33257 (AF129448); Salmonella enterica serovar Paratyphi BMX 85 18 241 (U77450); Salmonella enterica serovar Shomron BMX 74 02 019 (U77451); Salmonella enterica serovar Typhimurium CT18 (NC003198) and LT2 (NC003197 and X13854); Salmonella enterica subsp. arizonae serovar IIIa ATCC13314 (U77447); Salmonella enterica subsp. sofia BMX 84 01 519 (U77452); Serratia marcescens ATCC13880 (U77449); Shigella dysenteriae ATCC29027 (U77448); Shigella flexneri 2457T (NC004741); and Yersinia enterocolitica ATCC23715 (U77453).

28.3 Conclusions and Future Perspectives Molecular tests are a promising and necessary alternative for the identification of Klebsiella and Raoultella because several biochemical tests are currently needed to separate species, many species share a similar biochemical profile, and the presence of atypical strains, such as fastidious, slowgrowing or even pleiotropic, makes the identification even more difficult. rpoB gene sequencing has been shown to be a useful identification tool for different Enterobacteriaceae species, including Klebsiella. In addition, this technique can supplement incomplete or dubious biochemical identification. However, molecular tests for detection and quantification of Klebsiella from food are still necessary. Tests for food will have to cope with the issues of differing between environmental strains from strains with a pathogenic potential, if any differences exist, differing between a live or dead cell, and will probably have to be quantitative. Klebsiella is a repository of antimicrobial resistance genes.87 The more resistant strains are usually hospitalacquired, but multidrug-resistance is being already seen in the community. Persistent carries are an important source of resistance dissemination; however, the role of Klebsiella contaminated food in the spread of resistance genes must be

400

Molecular Detection of Foodborne Pathogens

S. dysenteriae S. flexneri 75 95 E. coli 1 70 E. coli 5 E. coli 2 68 Ec/Sh E. coli 4 92 E. coli 3 E. fergusonii K. ascorbata C. freundii E. cloacae E. aerogenes K. variicola k40 71 K. variicola k27 100 Kv ATCC BAA-830 77

99 72

P. mirabilis

P. stuartii

K. variicola k166 Kp ATCC 700603 Kp ATCC 13883

66 Kp k18 Kp MGH 78578 62 Kp ATCC 11296 Kp k14 55 73 Kp ATCC 13884 64 Rp k111 96 Rp k112 100 Ro ATCC 31898 Rp ATCC 33531 Rp/Ro Rt ATCC 33257 99 K. oxytoca k3 99 K. oxytoca k124 K. oxytoca k45 100 K. oxytoca k140 Ko 77 K. oxytoca k43 50 K. oxytoca k56 K. oxytoca ATCC 13182 96 99 K. oxytoca k50 76 K. oxytoca k89 Se BMX 84 01 519 100 99 Se BMX 74 02 019 Se ATCC 13314 Se Se CT18 57 Se BMX 85 18 241 96 70 Se LT2 (NC003197) Se LT2 (X13854) 71 H. alvei S. marcesecens Y. enterocolitica 63

76

Kv 96 Kp

98

S. saprophyticus

5%

Figure 28.2  Phylogenetic tree derived from partial 512 bp rpoB sequences of enterobacterial isolates. Numbers within the tree indicate the occurrence (%) of the branching order in 500 bootstrapped trees. Only values above 50 are shown. The scale bar indicates 5% divergence. Clusters are indicated by arrows. Cluster Ec/Sh: E. coli and Shigella spp.; cluster Kv: K. variicola; cluster Kp: K. pneumoniae; cluster Rp/Ro: R. planticola and R. ornithinolytica; cluster Ko: K. oxytoca; cluster Se: serovarsof S. enterica. Other enterobacterial species were not included in clusters. The rpoB sequence of Staphylococcus saprophyticus subsp. Saprophyticus strain ATCC 15305 (accession#NC007350) is included as outgroup.

addressed. Finally, an aspect to be solved is the importance of food as a source of more virulent K. pneumoniae strains, such as the hypermucoviscous serotype K1, and antibiotic

associated K. oxytoca colitis. A molecular test would be useful to detect and investigate the epidemiologic importance of such strains in food.

Klebsiella

Acknowledgments Work presented by the authors was supported by Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) and Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ) of Brazil, and the Fogarty International Program in Global Infectious Diseases (TW006563) of the National Institute of Health, which supported a postdoctoral fellowship for Rubens C.S. Dias.

References









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401 16. Granier, S.A. et al. New Klebsiella oxytoca β-lactamase genes blaOXY-3 and blaOXY-4 and a third genetic group of K. oxytoca based on blaOXY-3. Antimicrob. Agents Chemother., 47, 2922, 2003. 17. Podschun, R. et al. Incidence of Klebsiella species in surface waters and their expression of virulence factors. Appl. Environm. Microbiol., 67, 3325, 2001. 18. Abbott, S.L. Manual of Clinical Microbiology, 9th ed. ASM Press, Washington, DC, 2007, 698. 19. Munoz, M.A., et al. Molecular epidemiology of two Klebsiella pneumoniae mastitis outbreaks on a dairy farm in New York State. J. Clin. Microbiol., 45, 3964, 2007. 20. Struve, C. and Krogfelt, K.A. Pathogenic potential of environmental Klebsiella pneumoniae isolates. Environ. Microbiol., 6, 584, 2004. 21. Ko, W-C. et al. Community-acquired Klebsiella pneumoniae bacteremia: global differences in clinical patterns. Emerg. Infect. Dis., 8, 160, 2002. 22. Mandell, L.A. et al. Infectious disease society of America/ American thoracic society consensus guidelines on the management of community-acquired pneumonia in adults. Clin. Infect. Dis., 44, S27, 2007. 23. Keynan, Y. and Rubinstein, E. The changing face of Klebsiella pneumoniae infections in the community. Intern. J. Antimicrobial Agents, 30, 386, 2007. 24. Ni, Y-H. et al., Community-acquired brain abscess in Taiwan: etiology and probably source of infection. J. Microbiol. Immunol. Infect., 37, 231, 2004. 25. Fang, C-T. et al. Klebsiella pneumoniae genotype K1: an emerging pathogen that causes septic ocular or central nervous system complications from pyogenic liver abscess. Clin. Infect. Dis., 45, 284, 2007. 26. Turton, J.F. et al. Genetic similar isolates of Klebsiella pneumoniae serotype K1 causing liver abscess in three continents. J. Med. Microbiol., 56, 593, 2007. 27. Cheng, D.L. et al. Septic metastatic lesions of pyogenic liver abscess. Their association with Klebsiella pneumoniae bacteremia in diabetes patients. Arch Intern. Med., 151, 1557, 1991. 28. Habib, A.G. and Tambyah, P.A. Community-acquired Klebsiella pneumoniae central nervous system infections in adults in Singapore. Eur. J. Clin. Microbiol. Infect. Dis., 22, 486, 2003. 29. Chang, W.N. et al. Community-acquired spontaneous Klebsiella pneumoniae meningitis in adult cirrhotic patients with and without diabetes. Eur. J. Clin. Microbiol. Infect. Dis., 22, 271, 2003. 30. Andraca, R., Edson, R.S. and Kern, E.B. Rhinoscleroma: a growing concern in the United States? Mayo Clinic experience. Mayo Clin. Proc., 68, 1151, 1993. 31. De Champs, C. et al. Laryngeal scleroma associated with Klebsiella pneumoniae subsp. ozaenae. J. Clin. Microbiol., 43, 5811, 2005. 32. Chan, T.V. and Spiegel, J.H. Klebsiella rhinoscleromatis of the membranous nasal septum. J. Laryngol. Otol., 121, 998, 2007. 33. Medina, L.M. et al. Clinical, genetic and immunologic analysis of a family affected by ozena. Eur. Arch. Otorhinolaryngol., 260, 390, 2003. 34. Talmi, Y.P. et al. Computed tomography study of sinus involvement in ozena. Am. J. Rhinol., 9, 281, 1995. 35. Biedenbach, D.J., Moet, G.J. and Jones R.N. Occurrence and antimicrobial resistance pattern comparisons among bloodstream infection isolates from SENTRY Antimicrobial Surveillance Program (1997–2002). Diagn. Microbiol. Infect. Dis., 50, 59, 2004.

402 36. Podschun, R. et al. Isolation of Klebsiella planticola from newborns in a neonatal ward. J. Clin. Microbiol., 36, 2331, 1998. 37. Pessoa-Silva, C.L. et al. Extended-spectrum β-lactamaseproducing Klebsiella pneumoniae in a neonatal intensive care unit: risk factors for infection and colonization. J. Hosp. Infect., 53, 198, 2003. 38. Alves, M.S. et al. A case of severe pancreatitis complicated by Raoultella planticola infection. J. Med. Microbiol., 56, 696, 2007. 39. De Champs, C. et al. Prospective survey of colonization and infection caused by expanded-spectrum-β-lactamaseproducing members of the family Enterobacteriaceae in an intensive care unit. J. Clin. Microbiol., 27, 2887, 1989. 40. Peña, C. et al. Epidemiology and successful control of a large outbreak due to Klebsiella pneumoniae producing extendedspectrum β-lactamases. Antimicrob. Agents Chemother., 42, 53, 1998. 41. Asensio, A. et al. Outbreak of a multiresistant Klebsiella pneumoniae strain in an intensive care unit: antibiotic use as risk factor for colonization and infection. Clin. Infect. Dis., 30, 55, 2000. 42. Moustaoui, N. et al. Digestive tract colonization with extended spectrum beta-lactamase producing Enterobacteriaceae in a surgical intensive care unit in Casablanca. J. Hosp. Infect., 46, 238, 2000. 43. Berthelot, P. et al. Nosocomial colonization of premature babies with Klebsiella oxytoca: probable role of enteral feeding procedure in transmission and control of the outbreak with the use of gloves. Infect. Control. Hosp. Epidemiol., 22, 148, 2001. 44. Moustaoui, N. et al. Molecular biology of extended-spectrum β-lactamase-producing Enterobacteriaceae responsible for digestive tract colonization. J. Hosp. Infect., 57, 202, 2004. 45. De Almeida, V.C. et al. Genetic relatedness among extended-spectrum β-lactamase-producing Klebsiella pneumoniae outbreak isolates associated with colonization and invasive disease in a neonatal intensive care unit. Microb. Drug Resist., 11, 21, 2005. 46. Casewell, M.W. and Phillips, I. Aspects of plasmid-mediated antibiotic resistance and epidemiology of Klebsiella species. Am. J. Med., 70, 459, 1981. 47. Kristóf, K. et al. Extended-spectrum beta-lactamase-producing Klebsiella spp. in a neonatal intensive care unit: risk factors for the infection and the dynamics of the molecular epidemiology. Eur. J. Clin. Microbiol. Infect. Dis., 26, 563, 2007. 48. Lucet, J.C. et al. Control of a prolonged outbreak of extendedspectrum β-lactamase-producing Enterobacteriaceae in a university hospital. Clin. Infect. Dis., 29, 1411, 1999. 49. Cantón, R. et al. Prevalence and spread of extended-spectrum β-lactamase-producing Enterobacteriaceae in Europe. Clin. Microbiol. Infect., 14, 144, 2008. 50. Du, B. et al. Extend-spectrum beta-lactamase-producing Escherichia coli and Klebsiella pneumoniae bloodstream infections: risk factors and clinical outcome. Intensive Care Med., 28, 1718, 2002. 51. Bou, G. et al. Identification and broad dissemination of the CTX-M-14 β-lactamase in different Escherichia coli strains in northwest area of Spain. J. Clin. Microbiol., 40, 4030, 2002. 52. Colodner, R. Risk factors for the development of extendedspectrum beta-lactamase-producing bacteria in nonhospitalized patients. Eur. J. Clin. Microbiol. Infect. Dis., 23, 163, 2004.

Molecular Detection of Foodborne Pathogens 53. Pitout, J.D. et al. Emergence of Enterobacteriaceae producing extended-spectrum β-lactamases (ESBLs) in the community. J. Antimicrob. Chemother., 56, 52, 2005. 54. Blomberg, B. et al. High rate of fatal cases of pediatric septicemia caused by gram-negative bacteria with extendedspectrum beta-lactamases in Dar es Salaam, Tanzania. J. Clin. Microbiol., 43, 745, 2005. 55. Rennie, R.P. et al. Klebsiella pneumoniae gastroenteritis masked by Clostridium perfringens. J. Clin. Microbiol., 28, 216, 1990. 56. Guerrant, R L. et al. Role of toxigenic and invasive bacteria in acute diarrhea of childhood. N. Engl. J. Med., 293, 567, 1975. 57. Klipstein, F.A., and Engert R.F. Enterotoxigenic intestinal bacteria in tropical sprue. III. Preliminary characterization of Klebsiella pneumoniae enterotoxin. J. Infect. Dis., 132, 200, 1975. 58. Sabota, J.M. et al. A new variant of food poisoning: enteroinvasive Klebsiella pneumoniae and Escherichia coli sepsis from a contaminated hamburger. Am. J. Gastroenterol., 93, 118, 1998. 59. Favre-Bonte, S., Darfeuille-Michaud, A. and Forestier, C. Aggregative adherence of Klebsiella pneumoniae to human intestine-407 cells. Infect. Immun., 63, 1318, 1995. 60. Livrelli, V. et al. Adhesive properties and antibiotic resistance of Klebsiella, Enterobacter, and Serratia clinical isolates involved in nosocomial infections. J. Clin. Microbiol., 34, 1963, 1996. 61. Högenauer, C. et al. Klebsiella oxytoca as a causative organism of antibiotic-associated hemorrhagic colitis. N. Engl. J. Med., 355, 2418, 2006. 62. Tondo, E.C. et al. Identification of heat stable protease of Klebsiella oxytoca isolated from raw milk. Lett. Appl. Microbiol., 38, 146, 2004. 63. Liu, Y., et al. PCR detection of Klebsiella pneumoniae in infant formula based on 16S-23S internal transcribed spacer. Int. J. Food Microbiol., Epub Mar 20, 2008. 64. Muytjens, H.L. et al. Quality of powdered substitutes for breast milk with regard to members of the family Enterobacteriaceae. J. Clin. Microbiol., 26, 743, 1988. 65. Mollet, C., Drancourt, M. and Raoult, D. rpoB sequence analysis as a novel bases for bacterial identification. Mol. Microbiol., 26,1005, 1997. 66. Søgaard, M. et al. Peptide nucleic acid fluorescence in situ hybridization for rapid detection of Klebsiella pneumoniae from positive blood cultures. J. Med. Microbiol., 56, 914, 2007. 67. Kurupati, P. et al. Rapid detection of Klebsiella pneumoniae from blood culture bottles by real-time PCR. J. Clin. Microbiol., 42, 1337, 2004. 68. Mancini, N, et al. Molecular diagnosis of sepsis in neutropenic patients with haematological malignancies. J. Med. Microbiol., 57, 601, 2008. 69. Tenover, F.C. et al. Interpreting chromosomal DNA restriction patterns produced by pulsed-field gel electrophoresis: criteria for bacterial strain typing. J. Clin. Microbiol., 33, 2233, 1995. 70. Tenover, F.C., Arbeit, R.D. and Goering, R.V. How to select and interpret molecular strain typing methods for epidemiological studies of bacterial infections: a review for healthcare epidemiologists. Molecular Typing Working Group of the Society for Healthcare Epidemiology of America. Infect. Control. Hosp. Epidemiol., 18, 426, 1997.

Klebsiella 71. Gaillot, O. et al. Nosocomial outbreak of Klebsiella pneumoniae producing SHV-5 extended-spectrum β-lactamase, originating from a contaminated ultrasonography coupling gel. J. Clin. Microbiol., 36, 1357, 1998. 72. Gori, A. et al. Comparison of pulsed-field gel electrophoresis and randomly amplified DNA polymorphism analysis for typing extended-spectrum-β-lactamase-producing Klebsiella pneumoniae. J. Clin. Microbiol., 34, 2448, 1996. 73. Gazouli, M. et al. Study of an outbreak of cefoxitin-resistant Klebsiella pneumoniae in a general hospital. J. Clin. Microbiol., 35, 508, 1997. 74. Cartelle, M. et al. Risk factors for colonization and infection in a hospital outbreak caused by a strain of Klebsiella pneumoniae with reduced susceptibility to expanded-spectrum cephalosporins. J. Clin. Microbiol., 42, 4242, 2004. 75. Diancourt, L., et al. Multilocus sequence typing of Klebsiella pneumoniae nosocomial isolates. J. Clin. Microbiol., 43, 4178, 2005. 76. Paulin-Curlee, G.G. et al. Genetic diversity of mastitis-associated Klebsiella pneumoniae in dairy cows. J. Dairy Sci., 90, 3681, 2007. 77. Ko, K.S., et al. Clonal dissemination of extended-spectrumlactamase (ESBL)-producing Klebsiella pneumoniae isolates in a Korean hospital. J. Korean Med. Sci., 23, 53, 2008. 78. Hall, T.A., BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucl. Acids. Symp. Ser., 41, 95, 1999.

403 79. Food and Drug Administration. Investigations Operations Manual. FDA, Rockville, MD, 1993. 80. Downes, F.P. and Ito, K. Compendium of Methods for the Microbiological Examination of Foods, 4th ed. APHA, Washington, DC, 2001, chap.8. 81. Jiménez, F. et al. Evaluation of the presence of bacteria in food and environment of an Oncological Service of a National Hospital, San José, Costa Rica. Arch. Latinoam. Nutr., 54, 303, 2004. 82. Musgrove, M.T. et al. Enterobacteriaceae and related organisms isolated from shell eggs collected during commercial processing. Poult. Sci., 87, 1211, 2008. 83. Drancourt, M. and Raoult, D. Sequence-based identification of new bacteria: a proposition for creation of an orphan bacterium repository. J. Clin. Microbiol., 43, 4311, 2005. 84. Tamura, K. et al. Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Mol. Biol. Evol., 24, 1596, 2007. 85. Kimura, M., A simple method for estimating evolutionary rate of base substitutions through comparative studies of nucleotide sequences. J. Mol. Evol., 16, 111, 1980. 86. Saitou, N. and Nei, M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol. Biol. Evol., 4, 406, 1987. 87. Bistué, A.J.C.S., et al. Klebsiella pneumoniae multiresistance plasmid pMET1: similarity with the Yersinia pestis plasmid pCRY and integrative conjugative elements. Proc. Natl. Acad. Sci., 3, e1800, 2008.

29 Plesiomonas

Jesús A. Santos, Andrés Otero, and María-Luisa García-López University of León

Contents 29.1 Introduction.................................................................................................................................................................... 405 29.1.1 History and Current Taxonomic Status............................................................................................................ 405 29.1.2 Habitat............................................................................................................................................................... 406 29.1.3 Factors Affecting P. shigelloides in Foods....................................................................................................... 406 29.1.4 Characteristics of Disease................................................................................................................................. 407 29.1.4.1 Gastroenteritis.................................................................................................................................. 407 29.1.4.2 Extraintestinal Disease.................................................................................................................... 407 29.1.5 Pathogenicity and Virulence Factors................................................................................................................ 408 29.1.5.1 Toxins............................................................................................................................................... 408 29.1.5.2 Invasiveness..................................................................................................................................... 408 29.1.5.3 Hemolytic Activity........................................................................................................................... 409 29.1.5.4 Plasmids........................................................................................................................................... 409 29.1.5.5 Other Putative Virulence Factors.................................................................................................... 409 29.1.6 Control.............................................................................................................................................................. 409 29.1.7 Laboratory Diagnosis of P. shigelloides........................................................................................................... 409 29.1.7.1 Conventional Diagnosis................................................................................................................... 409 29.1.7.2 Molecular Diagnosis of P. shigelloides............................................................................................410 29.1.7.3 Typing...............................................................................................................................................411 29.2 Methods...........................................................................................................................................................................411 29.2.1 Reagents and Equipment....................................................................................................................................411 29.2.1.1 Reagents............................................................................................................................................411 29.2.1.2 Equipment and Supplies...................................................................................................................411 29.2.2 Sample Collection and Preparation...................................................................................................................412 29.2.3 PCR Assay and Visualization............................................................................................................................412 29.3 Conclusions and Further Perspectives.............................................................................................................................413 Acknowledgments.......................................................................................................................................................................413 References...................................................................................................................................................................................413

29.1  Introduction The genus Plesiomonas (plesios, neighbor, monas, unit, for neighbor to Aeromonas) consists of only one homogeneous species with a single DNA hybridization group (P. shigelloides) which comprises motile, facultative anaerobic, oxidase- and catalase-positive, glucose-fermenting, Gramnegative rod shaped bacteria that can cause serious gastroenteritis and extraintestinal infections in humans, particularly in immunocompromised hosts. This introduction provides a general overview of what is currently known about P. shigelloides in foods.

29.1.1 History and Current Taxonomic Status The long and complex history of the taxonomy of this genus has been reviewed by several authors.1–3 Briefly, the bacterium now known as P. shigelloides was first isolated in 1947 from

the feces of a patient and designated Strain C27. Ferguson and Henderson4 described this culture as “a motile ­organism containing a somatic antigen similar to the major antigen of Shigella sonnei phase 1.” At that time Strain C27 was thought to be a member of the family Enterobacteriaceae that “appeared to be an anaerogenic paracolon.” Later it was suggested that Strain C27 and S. sonnei might be put in the genus Esherichia as a new species “Escherichia sonnei.” In 1954, because of the cytochrome oxidase activity and lophothrichous flagella, the C27 organism was proposed to be moved to the genus Pseudomonas with the species name of shigelloides, the species epithet resulting from its Shigella-like characteristics. The name Pseudomonas michigani was also proposed because C27 was first isolated in Michigan. In 1961, the organism was transferred to the genus Aeromonas based upon its fermentative metabolism, flagellar arrangement and biochemical properties. The transfer of C27 strain to the generic 405

406

name Plesiomonas was proposed in 1962 since the organism did not exhibit some important characters, such as the enzymatic activity, traditionally linked to the genus Aeromonas. One year later, there was a new proposal to transfer C27 to a separated genus, Fergusonia, but the name P. shigelloides was confirmed because it was the first validly published. For over two decades, Plesiomonas has remained within the family Vibrionaceae until, based on genetic data; it was transferred to the family Enterobacteriaceae, this genus being the only oxidase-positive member of this family. Some authors5 had suggested that P. shigelloides could belong to a new taxonomic family, Plesiomonadaceae, and others2 have suggested that the appropriate family or nearest neighbours to the genus Plesiomonas may not have been discovered yet. Recently, using multilocus sequencing typing (MLST), Salerno et al.6 precisely defined the phylogenetic position of P. shigelloides within the family Enterobacteriaceae. The authors reported that the concatened sequence of four proteinencoding genes places P. shigelloides on a unique branch within the phylogenetic diversity of Enterobacteriaceae and concluded that: (i) in contrast to suggestions based on the 16S rRNA gene sequence, P. shigelloides should not be considered an atypical early branching member of this family, and (ii) the oxidase activity appears to be a derived characteristic that evolved secondarily from an oxidase-negative ancestor. Furthermore, their data ruled out a proposal, based on 5S rRNA, linking P. shigelloides with Proteus species.

29.1.2 Habitat P. shigelloides is primarily an aquatic organism. It is usually found in fresh surface water (rivers, lakes, streams, ponds, and sediments) or estuarine water, but can also be recovered from marine environments.1,2,7 Its minimum growth temperature (8°C) influences the more frequent occurrence in tropical and subtropical climates and also the seasonal incidence in temperate climates. It should be noted, however, that P. shigelloides has been recovered from aquatic environments in cold geographical areas such as Central Europe, Sweden, and even from a lake situated north of the Polar Circle.8 Medema and Schets9 have also reported that Plesiomonas recovery and density significantly correlated with index parameters for the trophic state (Secchi depth, a measurement of water clarity, and chlorophyll A) and for fecal pollution (Escherichia coli). Thus, in addition to temperature, the availability of nutrients and the level of sewage pollution are factors that support increasing concentrations of P. shigelloides in surface water. The organism has also been isolated from a wide range of warm and cold-blooded species other than man. Fish and shellfish in the natural habitats of P. shigelloides appear to act as secondary reservoirs for the bacterium, which has been isolated from crabs, bivalve molluscs, and from the intestines of 59% of freshwater fish in Zaire, from 10.2% of freshwater fish in Japan and from 23% of marine fish in Spain.7,10–12 Wild birds have also been found to carry P. shigelloides and most bird species from which the bacterium has been isolated either live in aquatic ecosystems and/or feed on fish.7 Isolation of

Molecular Detection of Foodborne Pathogens

P. shigelloides has also been reported from marine mammals, reptiles, and amphibians, monkeys, food animals (pigs, cattle, poultry, sheep, and goats) and pet ­animals (cats and dogs). The organism has been associated with disease in cats, freshwater fish, reptiles, and turtles. An extensive review of P. shigelloides from the veterinary perspective is that of Jagger.7 P. shigelloides is not a part of the normal gut flora of man and the rate of carriage among healthy people is generally low (0.2–3.2%) although it varies considerably with location, the highest rates corresponding to tropical and subtropical countries.

29.1.3  Factors Affecting P. shigelloides in Foods Most human P. shigelloides infections are suspected to be waterborne. Untreated or poorly chlorinated drinking water has been implicated in a number of large outbreaks of gastroenteritis with high attack rates.13,14 In addition, a number of cases have been associated with recreational use of water15 and water from aquaria.16 Snake-to-human transmission of P. shigelloides gastrointestinal infection has also been reported.17 Outbreaks of Plesiomonas-associated gastroenteritis have been attributed to contaminated oysters and also to fish (salt mackerel and cuttlefish salad), crab, shrimp, scallops, and shushi consumption.1,2,18 In the USA, infection with P. shigelloides has been strongly associated with eating raw or undercooked shellfish, usually raw oysters, and with travelling to high-risk areas (i.e., Mexico or Southeast Asia). The high incidence in Japan has been linked with dietary habits and also with trips to other Asian countries.19 Thus, the usual route of transmission of the organism in sporadic or epidemic cases is by ingestion of contaminated water or raw shellfish. In fact, there is only one known case associated with food other than water or fish, chicken being implicated.20 The bacterium also poses a hazard for people with water-related occupations or interested in water-related sports. Although information on the effects of intrinsic and extrinsic factors on the growth of P. shigelloides is limited to a few papers, a review on this subject has been undertaken by the International Commission on Microbiological Specifications for Foods.21 The minimum growth temperature is widely accepted to be 8ºC although at least one strain has been reported to grow at 0ºC.22 Optimum growth appears to occur in the range 37–39ºC and the maximum growth temperature is around 45ºC. For isolates from fish and clinical specimens, Herrera23 found that none grew at 6 or 8ºC, but most did at 10ºC although it required at least 9 days for visible growth to occur. Strains of this organism can be recovered from frozen foods stored by years. Temperatures of 42–44ºC have been recommended for the isolation of P. shigelloides when the presence of other organisms poses a problem.24 Miller and Koburger25 characterized 40 P. shigelloides strains from a variety of sources (water, sediment, fish, bivalve molluscs, and stools) and concluded that the organism is able to withstand up to 5% NaCl in trypticase soy broth but only

Plesiomonas

up to 3% in a less nutritious medium such as tryptone broth. All cultures grew in the pH range 4.5–8.5, with 60% growing at pH 4 and 85% at pH 9. Salt tolerance of the isolates tested in trypticase soy broth by Herrera23 was dependent on strain origin; thus strains from clinical specimens tolerated less salt (3.5%) than those from marine environments (5.5%). For all isolates, tolerance to acid and alkaline conditions was similar to that reported by Miller and Koburger.25 Even fewer studies on P. shigelloides growth under reduced water activity (aw) conditions have been reported. Minimum aw for growth of P. shigelloides, which varied with strain, and the type of humectant used, ranged from 0.92 (glycerol) to 0.97 (NaCl and sucrose).23 Knowledge of the effect of food processing on P. shigelloides is also extremely limited. Studies to date indicate that pasteurization at 60ºC for 30 min is an effective means of destroying P. shigelloides cells.25 Ingham26 evaluated the effects of storage in air and under vacuum and a modified atmosphere, consisting of 80% CO2, on the growth of P. shigelloides spiked on cooked crayfish tails. The organism did not grow at 8ºC under any packaging system. At 11ºC, the organism grew well in air but was strongly inhibited by the modified atmosphere and, to a lesser extent, by vacuum, and at 14ºC, only the modified atmosphere inhibited P. shigelloides growth. As for other Gram-negative, mesophilic, facultative anaerobic pathogenic bacteria, low temperatures and CO2 are effective means in preventing P. shigelloides growth in raw foods packaged under oxygen reduced conditions.

29.1.4 Characteristics of Disease Human infections caused by P. shigelloides can be divided into two major groups: gastroenteritis and extraintestinal disease. 29.1.4.1 Gastroenteritis Gastroenteritis is the most common illness associated with P. shigelloides infection.27 There are a number of epidemic outbreaks of diarrhea attributed to P. shigelloides as the causative agent and a series of case reports predominantly from tropical and subtropical countries. In cold and temperate climates, Plesiomonas cases of gastroenteritis are often associated with foreign travel. Traveller’s diarrhea is seen year-round whereas locally acquired infections show a marked seasonal trend related to environmental contamination of freshwater, with the peak occurring during the warmer seasons.19,28,29 Approximately 70% of people who present diarrhea linked to this organism have either an underlying disease (cancer, Crohn’s disease, achlorhydria, diverticulosis, cirrhosis, etc.) or an identifiable risk factor (e.g., tropical travel and/or seafood consumption).2,30 Several authors28,31 have reported that in contrast with other pathogenic foodborne bacteria such as Aeromonas or Yersinia enterocolitica, the organism affects all age groups although most studies indicate a higher incidence in adults. Both males and females are equally affected. At least three major clinical presentations of P. shigelloides gastroenteritis occur: a secretory (watery) type of

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diarrhea, a more invasive disease resembling shigellosis and a subacute or chronic disease lasting between 2 weeks and 2–3 months.27,31 On rare occasions, the organism, in association with other enteric pathogens (i.e., Aeromonas sobria), has been reported to cause a cholera-like disease.32 Brenden et al.31 concluded that Plesiomonas diarrhea is most often secretory in nature, but findings from Holmberg et al.30 and Kain and Kelly28 suggest that the dysenteric form (enteroinvasiveness) rather than the secretory (enterotoxin production) diarrhea is the most common presentation of Plesiomonas gastroenteritis. The infective dose is unknown but is presumed to be very high.2 On average, symptoms may begin between 24 and 50 h after consumption of contaminated food or water although shorter incubation times of 1–1.5 h have been reported.33 Diarrhea is the predominant symptom occurring in 94% of cases. Accompanying symptoms vary, but severe abdominal pain or cramping, nausea and/or vomiting and low grade fever, are most common. Less frequent symptoms include chills, headache, and some degree of dehydration.1,2,28,30 Patients with stool positive for P. shigelloides have either watery diarrhea or diarrhea with blood and mucus. The secretory form varies in severity from a mild illness of short duration, to severe diarrhea. It is usually reported to last between 1 and 7 days but can persist as long as 21 days, with up to 30 stools per day at the peak of the disease. The dysenteric form is usually severe, being characterized primarily by abdominal pain with the presence of macroscopic blood and mucus in the stools, which are often greenish tinged and slimy.31 In most cases of P. shigelloides gastroenteritis, the episode resolves spontaneously in a few days but in a significant proportion, infection becomes sub acute or chronic and does not resolve until the patient is placed on appropriate antimicrobial therapy. There have also been reports of fatal cases subsequent to primary enteric illness.2,32,34 For P. shigelloides diarrhea, the drugs of choice have been tetracycline or trimethoprim-sulfamethoxazole. It should be noted that P. shigelloides strains present natural resistance to penicillins, roxithromycin, clarithromycin, lincosamides, streptogramins, glycopeptides, and fusidic acid.23,35 29.1.4.2 Extraintestinal Disease Extraintestinal infections due to P. shigelloides are relatively rare. The organism has been implicated in cases of septicemia, meningitis, intra-abdominal infections (cholecystitis, pseudoappendicitis, peritonitis), endophtalmitis and other ocular diseases, wound infections, cellulitis, osteomyelitis, arthritis, proctitis, and pyosalpingitis. Most episodes of septicemia occur in adults, with the gastrointestinal tract being the source although some may originate from infected wounds or trauma. Meningitis is usually associated with neonates and appears to be vertically transmitted during birth. In the case of neonatal meningitis, the fatality rate approach 80%. Although predisposing factors leading to disseminated P. shigelloides infections are at present poorly defined, people with conditions leading to an impaired immune function are thought to be at increased risk of developing systemic disease.2

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29.1.5  Pathogenicity and Virulence Factors Although there has been interest in the pathogenicity of Plesiomonas, many aspects remain unknown. One of the problems is the lack of animal models for conclusive identification of virulence determinants and the negative result obtained in one human volunteer study.36 More recently, Vitovec et al.37 studying co-infection with P. shigelloides, Aeromonas spp. and Cryptosporidium parvum indicated that experimental infection of neonatal BALB/c mice, may serve as an initial experimental animal model for studying the initial steps of gastrointestinal colonization and the diarrheal disease syndrome caused by these bacterial and protozoan pathogens and that such a model can serve as a step leading toward improved models for understanding gastrointestinal colonization and the diarrheal disease syndrome. Current evidence indicates that the exact mechanism of P. shigelloides pathogenicity is not fully elucidated and that more than one virulence factor is required to cause diarrhea. Experience with other enteric pathogens (e.g., mesophilic Aeromonas, Yersinia enterocolitica or Escherichia coli) suggests that perhaps only some strains can cause disease in certain host population. A number of putative virulence factors have been identified, but none of them is widely accepted as being important for Plesiomonas-associated infections or tested for routinely. In addition, very few have been investigated in any detail. 29.1.5.1 Toxins Whether P. shigelloides produce toxins or not is a controversial issue. Sanyal et al.38 screened six P. shigelloides isolates for enterotoxigenicity in the adult rabbit ligated ileal loop assay. Very few positive (four out of 29) results were observed, with some of the strains producing only a small amount of fluid accumulation within the intestinal lumen. Ljungh et al.39 reported a positive Y-1 adrenal cell assay but a negative rabbit ileal loop assay with two strains of P. shigelloides. Gurwith and Williams40 assayed two strains of P. shigelloides by means of the Y-1 adrenal cell assay and both strains elicited a positive reaction for enterotoxin. Penn et al.41 reported that heat-labile and heat-stable enterotoxins were negative on the P. shigelloides strains isolated from a patient. Negative results for the production of enterotoxin and cytotoxin were also reported by Johnson and Lior42 and by Pitarangsi et al.43 Saraswhati et al.44 described the production of heat stable and heat labile enterotoxins from all 70 Plesiomonas strains tested, irrespective of origin or serotype. Partial purification and characterization of the heat-labile and heat-stable enterotoxins by Manorama et al.45 revealed that the former was active in the rabbit ileal loop and the latter in both the rabbit ileal loop and suckling mice. Gardner et al.46 grew 28 strains of P. shigelloides and a type strain in an iron-depleted medium. Filtrates of 24 of the 29 strains produced elongation of CHO cells. These changes could be prevented by heating or by preincubation of the filtrate culture with cholera antitoxin, the authors concluding that P. shigelloides elaborates a cholera-like toxin. Later on, they demonstrated complete inhibition of production of this CHO cell

Molecular Detection of Foodborne Pathogens

e­ longation factor(s) by growth in an iron-containing medium.47 Herrington et al.36 reported that genetic probes for heat-stable enterotoxins related to those of enterotoxigenic Escherichia coli were negative and also that heat-labile enterotoxins were not detected when a modified GM1-enzyme linked immunosorbent assay was used. According to Matthews et al.,48 the key to detecting Plesiomonas enterotoxins may be serial passage of isolates in vivo, also suggesting that the enterotoxin of P. shigelloides appears to be novel. The fact that enterotoxic activity appears to be rapidly lost upon in vitro subculture could explain the failure of many early studies to consistently identify such a factor.27 Falcón et al.49 reported that two out of five P. shigelloides strains from water were positive for enterotoxin production when tested in the suckling mouse model. They also observed cytotoxic activity in CHO, HeLa, and Vero cells, which was lost when the cell-free supernatants were heated at 56ºC for 15 min and partially inactivated by antiserum to the cytotoxin of Aeromonas hydrophila. Okawa et al.50 found and isolated a new heat-stable cytotoxin (anticholera toxin-reactive protein-lypopolysaccharide complex, ACRP-LPS complex) from the culture filtrate of one clinical P. shigelloides strain, which also gave a positive reaction in the suckling mouse model and suggested that both cytotoxicity and enterotoxigenicity by ACPR in the cytotoxin was the most important virulence factor in the pathogenesis of their P. shigelloides strain. 29.1.5.2  Invasiveness Information on P. shigelloides mechanisms involved in attachment, chemotaxis, and penetration of the gastrointestinal epithelium and its associated mucous layer is scarce. Adherence to host cells is a fundamental step in bacterial infection and many enteropathogens possess surface structures that facilitate adherence to host cell epithelial surfaces.51 For P. shigelloides, it is not known if flagella have a role in adherence. A glycocalyx was not detected on any P. shigelloides isolates by Theodoropoulos et al.52 although there is one report of a glycocalyx detected on the outer surface of this bacterium.18 Tsugawa et al.53 demonstrated that GroEL, known to be a chaperone or heat-shock protein, has a positive influence on the attachment of P. shigelloides to Caco-2 cells. The Sereny test has always yielded uniformly negative results.18,31 Studies investigating the invasive capability of P. shigelloides by using cell cultures (namely, Y1 mouse adrenal tumor cells, Hep-2 derived from larynx human carcinoma and HeLa derived from human cervical carcinoma) have been inconclusive since some of them obtained positive results54 while others did not find invasive potential.36,55,56 By means of transmission electron microscope, Theodoropoulos et al.52 demonstrated that isolates of P. shigelloides derived from clinical samples and one type strain were capable of adhering to and entering the human colon carcinoma cell line Caco-2. Their observations suggested that initial uptake may occur through a phagocytic–like process and also that noted free P. shigelloides in the cytosol of Caco-2 cells was due to escape from cytoplasmic vacuoles. The inconclusive data from previous studies were attributed to the no enteric cell lines chosen,

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Plesiomonas

which are not representative of the target cells in vivo. The authors also indicated the possibility that, like E. coli, P. shigelloides comprises different pathogenic phenotypes. Based on an invasion assay and flow cytrometry, the mechanisms of invasion of the P. shigelloides clinical isolate strain P-1 into Caco-2 cells have been studied by Tsugawa et al.57 The authors suggested that the P-1 strain invades through signal transduction followed by the rearrangement of the cytoskeletal proteins, which is important for the endocytosis of bacteria by the epithelial cells. They also conclude that the invasion of P. shigelloides induces apoptotic cell death. Finally, studying the interplay between this bacterium and host defence mechanisms, Pavlova et al.58 presented an evidence of the inhibitory activities of P. shigelloides toward papain-like proteinases and proposed that cathepsin inhibition is of importance to the survival and spread of this pathogen in a mammalian host. 29.1.5.3 Hemolytic Activity Efficient mechanisms for iron acquisition from the host during an infection are considered essential for virulence. A mechanism for iron acquisition is the production of hemolysins, which release iron from intracellular heme and hemoglobin. For P. shigelloides, the detection of hemolytic activity has also been questionable, but using appropriate methods, a number of workers have provided evidence that most isolates from different sources and geographic areas have the ability to display cytolytic activity against erythrocytes.49,59–63 In addition, the genes encoding the heme iron utilization system of P. shigelloides, which is similar to that of Vibrio cholerae, have been isolated and characterized. One of them, the hugA gene, encodes an outer membrane receptor, HugA, required for P. shigelloides heme iron utilization.64 Detection of the hugA gene is common in clinical and environmental P. shigelloides isolates.12 The presence of hemolytic molecules and the HugA outer membrane receptor may represent an important pathway for iron acquisition. Furthermore, vacuolating activity associated to hemolysin production has been reported for some strains of P. shigelloides.49 The hemolysin(s) roles in the ability to escape from the intravacuolar compartment and enterotoxigenicity have also been suggested.49,52 In conclusion, the above studies indicate that P. shigelloides could produce, at least, two hemolytic factors, their expression and detection being influenced considerably by environmental growth conditions and testing procedures.49,59,60,62 For this bacterium, the overlay assay appears to be the best routine procedure for detecting hemolytic activity. 29.1.5.4  Plasmids Many P. shigelloides strains contain plasmids. A very large plasmid (>150 MDa, ca. 230 kb) was found by Holmberg et al.30 in 12 of 27 clinical isolates, but it was not the same as the large virulence plasmid described for Shigella species and enteroinvasive Escherichia coli. The five clinical isolates studied by Herrington et al.36 also possessed in common a very large plasmid (between 118 and 312 MDa), which in gnotobiotic piglets appeared to facilitate the uptake of P. shigelloides into the mucosa of the distal ileum. Furthermore, when one

gnotobiotic piglet was infected with a cured strain, the animal remained healthy. It is possible that unstable virulence plasmids may be involved in P. shigelloides pathogenicity. Another plasmid-mediated property is antimicrobials resistance. Marshall et al.65 examined five P. shigelloides strains isolated from Louisiana blue crabs (Callinectes sapidus) for resistance to selected antibiotics and presence of plasmids. They found that each isolate carried three plasmids of approximately 2.5 kb, 3.8 kb, and 5.3 kb. Plasmid curing linked the streptomycin resistance determinant with the 3.8 kb and/or 5.3 kb plasmids. However, others workers have failed in demonstrating any association between plasmids and antibiotic resistance.66 29.1.5.5 Other Putative Virulence Factors P. shigelloides strains show elastolytic activity, which may be involved in connective tissue degradation. Santos et al.61 reported that elastin degradation was cell-associated, enhanced when the strains where grown in an iron-depleted medium and lost after thermal treatment at 100ºC for 10 min. They also found that the enzymatic activity was inactivated by phenyl methyl sulfonyl fluoride that is an inhibitor of serine proteases. Tetrodotoxin (TTX) is a potent marine neurotoxin, named after the order of fish from which it is most commonly associated, the Tetraodontiformes. The toxin can be produced by different bacterial species, one of them being P. shigelloides.67,68 This bacterium has also been identified as a histamine producer in scombroid fish species by LópezSabater et al.69 and González-Rey.3 The former authors suggested that P. shigelloides could play an important role within fish histidine decarboxylating bacteria because of its association with aquatic environments.

29.1.6 Control As the usual route of transmission of the organism in sporadic or epidemic cases is by ingestion of contaminated water and raw or undercooked fish and shellfish, the risk of infection can be reduced by avoiding the use of untreated water for drinking and food preparation, by maintaining appropriate heating temperatures for fishery products and by avoiding contamination of cooked or processed foods. Public health measures such as routine testing of drinking water for microbial indicators and free chlorine substantially also reduce waterborne outbreaks and subsequent morbidity. Appropriate chill storage, salting conditions and CO2 will prevent growth of the organism.

29.1.7 Laboratory Diagnosis of P. shigelloides 29.1.7.1  Conventional Diagnosis P. shigelloides can be found on several kinds of samples (clinical and environmental, including water, fishes, and foods) and the purpose of the analysis can be very different on each situation. The selection of a concrete procedure of analysis will be influenced by: (i) the required selectivity

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(mainly, the ability to suppress background microbiota); (ii) the desired diagnostic feature (mainly, the ability to differentiate between P. shigelloides and other morphologically or biochemically similar bacteria); (iii) the presumed presence of injured cells; and (iv) the need for quantifying the Plesiomonas population. The classical procedure for detection P. shigelloides on clinical and environmental samples includes the isolation of more or less typical colonies on a selective and differential agar and the confirmation of the identity of isolates by a set of morphological and biochemical tests. For the isolation, different solid media have been used, particularly enteric agars (MacConkey agar, salmonella–shigella agar, xylose lysine deoxycholate agar, Hektoen enteric agar, deoxycholate lactose agar and Endo agar, among others). However, the morphology of Plesiomonas colonies on such media is very similar to that of colonies of other growing bacteria (Aeromonas, Enterobacteriaceae). Two differential media, formulated specifically for the isolation and enumeration of P. shigelloides are inositol brilliant green bile salts agar (IBB)70 and Plesiomonas agar (PL).1 Besides the selective components (brilliant green and bile salts) IBB agar includes inositol as carbon source, which can be used only for a few competing bacteria. At the same time, Plesiomonas ferments inositol, giving red to pinkish colonies on IBB, while nonfermeting inositol strains, such as Aeromonas strains, appear colourless (in over-crowded plates, Plesiomonas colonies can appear small and white)71 and coliforms form greenish or pink colonies.72 On PL agar lysine decarboxylase-positive and nonfermenting organisms such as P. shigelloides form pink colonies surrounded by a red zone.73 However, because of the reduced concentration of bile salts, PL agar is less inhibitory of the competitive microbiota than IBB agar, the latter being generally preferred for the isolation of Plesiomonas from environmental samples. On the other hand, PL agar shows a better performance in recovering heat and cold stressed cells of Plesiomonas.73 For routine analysis of environmental samples incubation of plates at 35ºC for 24–48 h is generally recommended,74 though 44ºC was found to be the optimal incubation temperature to easily differentiate colonies of Plesiomonas and Aeromonas in 24 h when Plesiomonas differential agar is used.75 Enrichment of samples before plating is a controversial practice. Four media have been proposed: alkaline peptone water (APW), bile peptone broth, and tetrathionate broth with or without iodine. As an example of the contradictory results, Freund et al.76 found that enrichment with tetrathionate broth without iodine gave a high recovery rate for P. shigelloides from fresh water, whereas Van Damme and Vandepitte11 reported no increase in isolation rate of P. shigelloides from freshwater fish using tetrathionate, with or without iodine. If tetrathionate broth (with or without iodine) is selected as enrichment media, incubation at 40ºC for 24 h seems to provide the highest recovery of P. shigelloides.76 In order to increase the recovery of P. shigelloides, it is a ­common practice to combine the direct plating on two selective

Molecular Detection of Foodborne Pathogens

media (usually IBB agar and PL agar) and an enrichment on tetrathionate broth without iodine followed by streaking on the two selective media.24 The identification of suspect isolates presents little difficulties and it can be done by inoculation both in Triple Sugar Iron (TSI) slants and inositol gelatine deeps, with incubation at 35ºC for 24 h. An oxidase test and Gram stain are also necessary. P. shigelloides is a Gram-negative rod, oxidasepositive, alkaline over acid with no gas or hydrogen sulfide in TSI, ferments inositol and fails to hydrolyze gelatine. A complete list of useful characteristics of P. shigelloides can be found elsewhere.24 Alternatively to the classical biochemical identification, miniaturised systems, like API 20E,12 BBL Crystal E/NF,77 Phoenix 100 ID/AST and NID Panel78 offer reliable results. 29.1.7.2  Molecular Diagnosis of P. shigelloides Due to the low pathogenic potential of P. shigelloides, limited investigations have been conducted to facilitate the detection of the microorganism. The molecular methods reported for P. shigelloides are focused on PCR. A summary of the characteristics of the methods, including target, primers, visualization procedure and application is provided in Table 29.1. The first published procedure was directed toward specific sequences of the 23S rDNA,79 amplifying a fragment of 284 bp. This method was checked against a number of isolates of P. shigelloides from aquatic environments, clinical, and animal origin, but it was applied only to pure cultures and a procedure for the testing of foods commodities was not provided. Based on the PCR procedure of González-Rey et al.,79 Gu and Levin developed two modified methods that allowed for quantification of P. shigelloides.80,81 The first method was carried out in homogenates of shellfish tissue (clams and oysters) that were then seeded with known numbers of a collection strain of P. shigelloides.80 The bacteria were recovered by differential centrifugation and treated with formaldehyde. DNA was extracted by lysing the cells with a mixture of Triton X-100 and sodium azide and purified with a commercial clean-up system. After PCR amplification, the PCR products were resolved by electrophoresis in controlled conditions and stained with GelStarTM (Cambrex, Rockland, MN). The gel was photographed and the fluorescent intensities of the DNA band were analyzed with the public domain Image software (http://rsb.info.nih.gov/ij/index.html). The relative fluorescent intensity of the PCR products was plotted against the log of CFU/g. The detection level was 60 CFU/g for clams and 200 CFU/g for oysters, but they were improved to 4 and 40 CFU/g respectively, with the introduction of a nonselective enrichment step. The second modification was developed to differentiate live and dead cells by using a DNA intercalating dye (ethidium bromide monoazide [EMA]) unable to penetrate into live cells.81 The dye would be incorporated mainly by dead cells with damaged membranes, inhibiting PCR amplification of the target DNA. At the same time, quantification of the viable cells is achieved by analyzing the relative fluorescent intensities of DNA bands obtained after a controlled PCR amplification.

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Plesiomonas

Table 29.1 Characteristics of PCR Assays Developed for P. shigelloides Target 23S rDNA 23S rDNA 23S rDNA

hugA

Primers and Probes (5′–3′)

Visualization

PS23FW3-CTC CGA ATA CCG TAG AGT GCT ATC C PS23RV3-CTC CCC TAG CCC AAT AAC ACC TAA A PS23FW3 PS23RV3 Forward-AGC GCC TCG GAC GAA CAC CTA Reverse-GTG TCT CCC GGA TAG CAC Probe LCRed640-GGT AGA GCA CTG TTA AGG CTA GGG GGT CAT C-P Forward-GCG AGC GGG AAG GGA AGA ACC Reverse-GTC GCC CCA AAC GCT AAC TCA TCA

Agarose gel electrophoresis (284 bp)

Pure cultures

González-Rey et al.79

Agarose gel electrophoresis (284 bp) Quantification by image analysis Real-Time FRET PCR

Clam and oyster tissue Stool samples

Gu and Levin80,81 Loh and Yap82

Agarose gel electrophoresis (435 bp)

Fish samples

Herrera et al.12

A quantitative assay was developed by Loh and Yap,82 based in the amplification of 23S rDNA, with the specific primers presented in Table 29.1. An LCRed640 labeled probe was also designed to specifically hybridizate to the amplicon. The reaction mixture included SYBR Green I, that acts as a generic donor in fluorescent resonance energy transfer (FRET) to excite LCRed640, and this is only possible if amplification occurs. The intensity of the measured fluorescence is proportional to the amount of DNA generated during the amplification process and the time of detection is related to the initial amount of DNA. The sensitivity of the assay was tested using serially diluted P. shigelloides DNA and the specificity was determined against 27 other bacterial species implicated in gastrointestinal disease. The PCR procedure was applied on stool samples from patients with diarrhea, using the QIAamp® DNA Stool Mini Kit (Qiagen, Hilden, Germany) for DNA extraction. A different PCR assay was developed by Herrera et al.12 for detection of P. shigelloides in fish samples. The PCR was directed towards the hugA gene, that encodes an outer membrane receptor required for heme iron utilization64 and probably implicated in the virulence mechanisms of Plesiomonas. The assay has been applied both to pure cultures of bacteria and to food samples. The selectivity was tested against strains of bacteria commonly found in aquatic environments or of relevance as foodborne pathogens. This procedure will be further detailed as a working protocol. 29.1.7.3 Typing Serotyping has been an important tool for differentiating among strains of P. shigelloides, since biotyping is of limited value, due to the phenotypic homogeneity of this species. Two major schemes were initially developed and later unified in an international antigenic scheme.83 Serotyping has been used for epidemiological studies with clinical and environmental strains,84,85 but it is now on the decline and is likely to be replaced by molecular methods. Even though many isolates of Plesiomonas carry plasmids (see Section 29.1.5.4), plasmid profiling does not seem a suitable procedure for epidemiological studies because of their heterogenicity. Molecular typing, as RAPD, PFGE and, more recently, MLST has been used to investigate the diversity

Application

Reference

and relationships of isolates from different origins.6,8,19,86 González-Rey et al.8 showed that RAPD and PFGE were able to discriminate among strains belonging to different serotypes, but not strains from the same serotype. All the methods tested had good performance; considering that RAPD is fast, simple, and inexpensive, it could be a promising method for routine typing of P. shigelloides until a standardized procedure is available.

29.2  Methods 29.2.1 Reagents and Equipment The reagents and equipment are listed without reference to any commercial brand. 29.2.1.1 Reagents • Tryptone soya broth plus 0.6% yeast extract (TSBYE) • Phosphate buffered saline (PBS) • Triton X-100 • Oligonucleotide primers: • Forward 5′-GCG AGC GGG AAG GGA AGA ACC-3′ • Reverse 5′-GTC GCC CCA AAC GCT AAC TCA TCA-3′ • PCR reagents: • 10× PCR buffer • 10 mM dNTP mixed solution • Recombinant thermostable DNA polymerase • Ultrapure double-distilled water • Agarose • Ethidium bromide • Electrophoresis loading buffer • Molecular weight marker (100 bp) 29.2.1.2 Equipment and Supplies • Micropipets and sterile filter-fitted tips • Microcentrifuge and microtubes • Incubator • Heating block • PCR tubes

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Molecular Detection of Foodborne Pathogens

• Thermal cycler • Electrophoresis system, including power supply, UV light and a gel documentation system

29.2.2 Sample Collection and Preparation P. shigelloides is considered a fishborne pathogen and the procedure that we have developed has been used successfully to test fish and seafood samples,12 but it may also work with other food samples. (1) Fish samples of 10 g are blended with 90 ml of TSBYE and incubated at 35±2ºC for 24 h with constant shaking (150 rev min–1). (2) One millilitre of the upper phase of the culture is withdrawn to a microcentrifuge tube and centrifuged for 5 min at 10000×g. The supernatant is carefully removed and the pellet is then processed for DNA extraction. If the procedure has to be interrupted, we have successfully frozen the pellets from the enrichment cultures in TSB plus 40% glycerol at –40ºC; before testing, 20 µl of the frozen culture is transferred to 5 ml of fresh TSBYE and incubated overnight at 37ºC. One millilitre of the culture is then centrifuged as described above. (3) The pellet is washed three times with PBS and resuspended in 500 µl of a 1% Triton X-100 solution. The suspension is boiled for 5 min and immediately cooled in ice. The boiled suspension is centrifuged at 10000×g for 5 min at 4ºC and the supernatant is transferred to a fresh tube and frozen at –20ºC for long-term storage. Other procedures can be used for DNA extraction from the washed pellet. In our laboratory we have checked commercial kits from different suppliers (GE Healthcare, http://www. gehealthcare.com; Qiagen, http://www.qiagen. com; Bio-Rad, http://www.biorad.com) with no differences in the performance of the PCR. (4) When the sample material is a pure culture of a microorganism, a suspension is prepared from a single colony in 200 µl of PBS and processed as described in step 3. (5) A similar enrichment procedure (TSB supplemented with yeast extract) was used for the detection of P. shigelloides in clam and oyster.80 The addition of formaldehyde at a final concentration of 4% was useful to decrease the inhibition of the PCR by enzymes present in the shellfish tissue. Moreover, the presence of 0.1% BSA to the PCR reaction mixtures improved the DNA amplification.

29.2.3  PCR Assay and Visualization (1) For each sample to be tested, make up a mix of 5 μl of 10× PCR buffer, 25 pmol of each primer (forward 5′-GCG AGC GGG AAG GGA AGA ACC-3′ and reverse 5′-GTC GCC CCA AAC GCT AAC









TCA TCA-3′), 0.1 mM of dNTPs, 0.8 μl (0.8 U) of DNA polymerase and 5 μl (ca. 50 ng) of template DNA, for a total reaction volume of 50 µl. A master mix can be prepared for all the amplification reactions of a PCR run. In our laboratory we routinely use enzymes and chemicals from Biotools, S.A (http://www.biotools.net) and from 5 Prime (http:// www.5prime.com) with good results. (2) Prepare a positive control with DNA from a reference strain of P. shigelloides (type strain ATCC 14029, supplied by the Spanish Type Culture Collection, CECT 4262, is the choice in our laboratory) and a negative control with 5 µl of double-distilled sterile water instead of DNA template. The controls are freshly prepared for every set of PCR reactions. (3) Briefly spin the tubes to ensure that the reaction mix is at the bottom of the tube. (4) Place tubes in a thermal cycler, fitted with a heated lid to prevent evaporation (we use a MasterCycler®, from Eppendorf, http://www.eppendorf.com), and run a program of an initial denaturation step at 94ºC for 3 min, followed by 30 cycles of denaturation at 92°C for 30 s, annealing at 63°C for 30 s, and extension at 72°C for 1.5 min. A final extension step is done at 72°C for 3 min. (5) Load 5 µl of the PCR products into a 1% (w/v) agarose gel and perform the electrophoresis until a good resolution of the bands is achieved (about 1 h at 6 V/cm). (6) Stain the gel with ethidium bromide and visualize under UV light.

Results obtained with different strains of P. shigelloides are shown in Figure 29.1. The presence of a band of 435 bp is considered a positive result.

1

2

3

4

5

6

Figure 29.1  Results of amplification of a 435 bp fragment of hugA gene from P. shigelloides. Lane 1, Molecular size marker 100 bp ladder; lane 2, P. shigelloides ATCC 14029 (type strain); Lane 3, P. shigelloides P1 (isolated from grouper); Lane 4, P. shigelloides P2 (isolated from halibut); Lane 5, P. shigelloides C1 (clinical strain isolated from a patient with diarrhea); Lane 6, negative control.

413

Plesiomonas

29.3 Conclusions and Further Perspectives Isolation of P. shigelloides from foods is not well defined. Several enrichment procedures and isolation media have been proposed, but there is evidence that none of them is particularly effective and a combination of enrichment broth incubated at 40ºC and isolation in two selective plating media is advisable to enhance the recovery of this bacterium.24,76 On the other hand, identification of the isolates presents little difficulty, either using conventional biochemical techniques or modern commercial miniaturised systems, like API 20E and BBL Crystal E/NF.12,77 Thus, the introduction of molecular methods is of great importance if they contribute to the detection and isolation of P. shigelloides and of minor interest for the identification of isolates. All the methods revised in this chapter work well for pure cultures, and two of them have been used to test food samples (Table 29.1). The alleged limitations of the procedures are related to the presence of PCR inhibitors in the food samples or in the enrichment broths and these limitations can be overcome by using nonselective enrichment broths, as TSB or TSBYE.12,80 The selectivity of the PCR assays for P. shigelloides has been checked against a number of strains of Plesiomonas isolated from different origins as well as strains of closely related bacteria, as members of the genera Vibrio, Aeromonas or Escherichia. Herrera et al.12 indicated a nonspecific amplification both with primers directed towards 23S rDNA and hugA gene with a particular strain of E. coli harbouring the stx2 gene, but it seems improbable that this situation would occur in the routine food analysis. Identification of suitable targets can be a promising way of develop new procedures for molecular detection of plesiomonads. In our laboratory we are exploring the potential use of the histidine decarboxilase gene, implicated in the production of histamine and characteristic of some Gram-negative bacteria, and the chiA gene, which codified for a chitinase presented in different species of marine bacteria. Also, the formulation of new culture media and the definition of new procedures to enhance the recovery of P. shigelloides from food samples will be of help. More molecular investigations are needed to elucidate the role of P. shigelloides in human and animal disease as well as to identify the pathogenic mechanisms and to facilitate the detection, identification, and characterization of this microorganism.

Acknowledgments The authors are grateful to the Spanish Ministry of Education and Science (grants AGL2002-762, AGL2004-4672/ALI and CONSOLIDER – Ingenio 2010: CARNISENUSA (CSD200700016)), the Junta de Castilla y León (grant LEO34A06 and research group GR155) and the University of León (grant ULE2001-12) for the financial support to study different aspects of Gram-negative foodborne pathogens.

References















1. Miller, M.A. and Koburger, J.A. Plesiomonas shigelloides: An opportunistic food and waterborne pathogen. J. Food Prot., 48, 449, 1985. 2. Janda, J.M. and Abbott, S.L. The genus Plesiomonas. In The Enterobacteria, 2nd ed. ASM Press, Washington, DC, 2006. 3. González-Rey, C. Studies on Plesiomonas shigelloides isolated from different environments. Ph.D. thesis, Department of Veterinary Microbiology, Swedish University of Agricultural Sciences, 2003. 4. Ferguson, W.W. and Henderson, N.D. Description of strain C27: a motile organism with the major antigen of Shigella sonnei phase I. J. Bacteriol., 54, 179, 1947. 5. Ruimy, R. et al. Phylogenetic analysis and assessment of the genera Vibrio, Photobacterium, Aeromonas, and Plesiomonas deduced from small-subunit rRNA sequences. Int. J. Syst. Bacteriol., 44, 416, 1994. 6. Salerno, A. et al. Recombining population structure of Plesiomonas shigelloides (Enterobacteriaceae) revealed by multilocus sequence typing. J. Bacteriol., 2007. 7. Jagger, T.D. Plesiomonas shigelloides: A veterinary perspective. Infect. Dis. Rev., 2, 199, 2000. 8. Gonzalez-Rey, C. et al. Unexpected finding of the “tropical” bacterial pathogen Plesiomonas shigelloides from lake water north of the Polar Circle. Polar Biol., 26, 495, 2003. 9. Medema, G. and Schets, C. Occurrence of Plesiomonas shigelloides in surface water: Relationship with faecal pollution and trophic state. Zbl. Hyg. Umweltmed., 194, 398, 1993. 10. Arai, T. et al. A survey of Plesiomonas shigelloides from aquatic environments, domestic animals, pets and humans. J. Hyg. (Lond), 84, 203, 1980. 11. Van Damme, L.R. and Vandepitte, J. Frequent isolation of Edwardsiella tarda and Plesiomonas shigelloides from healthy Zairese freshwater fish: A possible source of sporadic diarrhea in the tropics. Appl. Environ. Microbiol., 39, 475, 1980. 12. Herrera, F.C. et al. Occurrence of Plesiomonas shigelloides in displayed portions of saltwater fish determined by a PCR assay based on the hugA gene. Int. J. Food Microbiol., 108, 233, 2006. 13. Tsukamoto, T. et al. Two epidemics of diarrhoeal disease possibly caused by Plesiomonas shigelloides. J. Hyg. (Lond), 80, 275, 1978. 14. Centers for Disease Control and Prevention. Plesiomonas shigelloides and Salmonella serotype Hartford infections associated with a contaminated water supply—Livingston County, New York, 1996, MMWR, 47, 394, 1998. 15. Dziuban, E.J. et al. Surveillance for waterborne disease and outbreaks associated with recreational water-United States, 2003–2004. MMWR, 55, 1, 2006. 16. Centers for Disease Control and Prevention, Aquariumassociated Plesiomonas shigelloides infection—Missouri. MMWR, 38, 617, 1989. 17. Davis, W.A. et al. Snake-to-Human transmission of Aeromonas (Pl) shigelloides resulting in gastroenteritis. Southern Med. J., 71, 474, 1978. 18. Kirov, S.M. Aeromonas and Plesiomonas species. In Food Microbiology: Fundamentals and Frontiers, Doyle, M.P., Beuchat, L.R., and Montville, T.J., Eds. ASM Press, Washington, DC, 1997. 19. Shigematsu, M. et al. An epidemiological study of Plesiomonas shigelloides diarrhoea among Japanese travellers. Epidemiol. Infect., 125, 523, 2000.

414 20. Newsom, R. and Gallois, C. Diarrheal disease caused by Plesiomonas shigelloides. Clin. Microbiol. Newsl., 4, 158, 1982. 21. ICMSF. Microorganisms in Foods 5: Characteristics of Microbial Pathogens. Blackie Academic & Professional, London, 1996. 22. Rouf, M.A. and Rigney, M.M. Growth temperatures and temperature characteristics of Aeromonas. Appl. Microbiol., 22, 503, 1971. 23. Herrera, F.C. Microbiology of marine fish: indicator, spoilage and pathogenic microorganisms, with reference to Plesiomonas shigelloides. Ph.D. thesis, University of León, 2004. 24. Palumbo, S.A. et al. Aeromonas, Arcobacter, and Plesiomonas. In Compendium of Methods for the Microbiological Examination of Foods, Downes, F.P. and Ito, K., Eds., 4th ed. APHA, Washington, DC, 2001. 25. Miller, M.A. and Koburger, J.A. Tolerance of Plesiomonas shigelloides to pH, sodium chloride and temperature. J. Food Prot., 49, 877, 1986. 26. Ingham, S.C. Growth of Aeromonas hydrophila and Plesiomonas shigelloides on cooked crayfish tails during cold storage under air, vacuum and a modified atmosphere. J. Food Prot., 53, 665, 1990. 27. Clark, R.B. and Janda, J.M. Plesiomonas and human disease. Clin. Microbiol. Newsl., 13, 49, 1991. 28. Kain, K.C. and Kelly, M.T. Antimicrobial susceptibility of Plesiomonas shigelloides from patients with diarrhea. Antimicrob. Agents Chemother., 33, 1609, 1989. 29. Tseng, H.K. et al. Characteristics of Plesiomonas shigelloides infection in Taiwan. J. Microbiol. Immunol. Infect., 35, 47, 2002. 30. Holmberg, S.D. et al. Plesiomonas enteric infections in the United States. Ann. Intern. Med., 105, 690, 1986. 31. Brenden, R.A., Miller, M.A., and Janda, J.M. Clinical disease spectrum and pathogenic factors associated with Plesiomonas shigelloides infections in humans. Rev. Infect. Dis., 10, 303, 1988. 32. Sawle, G. et al. Fatal infection with Aeromonas sobria and Plesiomonas shigelloides. Br. Med. J., 292, 525, 1986. 33. Wouafo, M. et al. An acute foodborne outbreak due to Plesiomonas shigelloides in Yaounde, Cameroon. Foodborne Pathog. Dis., 3, 2006. 34. Nolte, F.S. et al. Proctitis and fatal septicemia caused by Plesiomonas shigelloides in a bisexual man. J. Clin. Microbiol., 26, 388, 1988. 35. Stock, I. and Wiedemann, B. Natural antimicrobial susceptibilities of Plesiomonas shigelloides strains. J. Antimicrob. Chemother., 48, 803, 2001. 36. Herrington, D.A. et al. In vitro and in vivo pathogenicity of Plesiomonas shigelloides. Infect. Immun., 55, 979, 1987. 37. Vitovec, J. et al. Enteropathogenicity of Plesiomonas shigelloides and Aeromonas spp. in experimental mono- and coinfection with Cryptosporidium parvum in the intestine of neonatal BALB/c mice. Comp. Immunol. Microbiol. Infect. Dis., 24, 39, 2001. 38. Sanyal, S.C., Singh, S.J., and Sen, P.C. Enteropathogenicity of Aeromonas hydrophila and Plesiomonas shigelloides. J. Med. Microbiol., 8, 195, 1975. 39. Ljungh, A., Popoff, M., and Wadstrom, T. Aeromonas hydrophila in acute diarrheal disease: detection of enterotoxin and biotyping of strains. J. Clin. Microbiol., 6, 96, 1977. 40. Gurwith, M.J. and Williams, T.W. Gastroenteritis in children: a two-year review in Manitoba. I. Etiology. J. Infect. Dis., 136, 239, 1977.

Molecular Detection of Foodborne Pathogens 41. Penn, R.G. et al. Plesiomonas shigelloides overgrowth in the small intestine. J. Clin. Microbiol., 15, 869, 1982. 42. Johnson, W.M. and Lior, H. Cytotoxicity and suckling mouse reactivity of Aeromonas hydrophila isolated from human sources. Can. J. Microbiol., 27, 1019, 1981. 43. Pitarangsi, C. et al. Enteropathogenicity of Aeromonas hydrophila and Plesiomonas shigelloides: prevalence among individuals with and without diarrhea in Thailand. Infect. Immun., 35, 666, 1982. 44. Saraswathi, B., Agarwal, R.K., and Sanyal, S.C. Further studies on enteropathogenicity of Plesiomonas shigelloides. Indian J. Med. Res., 78, 12, 1983. 45. Manorama, T.V., Agarwal, R., and Sanyal, S. Enterotoxins of Plesiomonas shigelloides: partial purification and characterization. Toxicon. Suppl., 3, 269, 1983. 46. Gardner, S.E., Fowlston, S.E., and George, W.L. In vitro production of cholera toxin-like activity by Plesiomonas shigelloides. J. Infect. Dis., 156, 720, 1987. 47. Gardner, S.E., Fowlston, S.E., and George, W.L. Effect of iron on production of a possible virulence factor by Plesiomonas shigelloides. J. Clin. Microbiol., 28, 811, 1990. 48. Matthews, B.G., Douglas, H., and Guiney, D.G. Production of a heat stable enterotoxin by Plesiomonas shigelloides. Microb. Pathog., 5, 207, 1988. 49. Falcón, R. et al. Intracellular vacuolation induced by culture filtrates of Plesiomonas shigelloides isolated from environmental sources. J. Appl. Microbiol., 95, 273, 2003. 50. Okawa, Y. et al. Isolation and characterization of a cytotoxin produced by Plesiomonas shigelloides P-1 strain. FEMS Microbiol. Lett., 239, 125, 2004. 51. Sansonetti, P.J. Bacterial pathogens, from adherence to invasion: comparative strategies. Med. Microbiol. Immunol., 182, 223, 1993. 52. Theodoropoulos, C. et al. Plesiomonas shigelloides enters polarized human intestinal Caco-2 cells in an in vitro model system. Infect. Immun., 69, 2260, 2001. 53. Tsugawa, H. et al. Cell adherence-promoted activity of Plesiomonas shigelloides GroEL. J. Med. Microbiol., 56, 23, 2007. 54. Binns, M.M. et al. Invasive ability of Plesiomonas shigelloides. Zbl. Bakt. Hyg. A, 257, 343, 1984. 55. Sanyal, S.C., Saraswathi, B., and Sharma, P. Enteropatho­ genicity of Plesiomonas shigelloides. J. Med. Microbiol., 13, 401, 1980. 56. Abbott, S.L., Kokka, R.P., and Janda, J.M. Laboratory investigations on the low pathogenic potential of Plesiomonas shigelloides. J. Clin. Microbiol., 29, 148, 1991. 57. Tsugawa, H. et al. Invasive phenotype and apoptosis induction of Plesiomonas shigelloides P-1 strain to Caco-2 cells. J. Appl. Microbiol., 99, 1435, 2005. 58. Pavlova, A. et al. Inhibition of mammalian cathepsins by Plesiomonas shigelloides. Folia Microbiol. (Praha), 51, 393, 2006. 59. Daskaleros, P.A., Stoebner, J.A., and Payne, S.M. Iron uptake in Plesiomonas shigelloides: cloning of the genes for the heme-iron uptake system. Infect. Immun., 59, 2706, 1991. 60. Janda, J.M. and Abbott, S.L. Expression of hemolytic activity by Plesiomonas shigelloides. J. Clin. Microbiol., 31, 1206, 1993. 61. Santos, J.A. et al. Hemolytic and elastolytic activities influenced by iron in Plesiomonas shigelloides. J. Food Prot., 62, 1475, 1999. 62. Baratéla, K.C. et al. Effects of medium composition, calcium, iron and oxygen on haemolysin production by Plesiomonas shigelloides isolated from water. J. Appl. Microbiol., 90, 482, 2001.

Plesiomonas 63. González-Rodríguez, M.N. et al. Cell-associated hemolytic activity in environmental strains of Plesiomonas shigelloides expressing cell-free, iron-influenced extracellular hemolysin. J. Food Prot., 70, 885, 2007. 64. Henderson, D.P. et al. Characterization of the Plesiomonas shigelloides genes encoding the heme iron utilization system. J. Bacteriol., 183, 2715, 2001. 65. Marshall, D.L., Kim, J.J., and Donnelly, S.P. Antimicrobial susceptibility and plasmid-mediated streptomycin resistance of Plesiomonas shigelloides isolated from blue crab. J. Appl. Bacteriol., 81, 195, 1996. 66. Kelly, M.T. and Kain, K.C. Biochemical characteristics and plasmids of clinical and environmental Plesiomonas shigelloides. Experientia, 47, 439, 1991. 67. Simidu, U. et al. Marine bacteria which produce tetrodotoxin. Appl. Environ. Microbiol., 53, 1714, 1987. 68. Cheng, C.A. et al. Microflora and tetrodotoxin-producing bacteria in a gastropod. Niotha clathrata, Food Chem. Toxicol., 33, 929, 1995. 69. Lopez-Sabater, E.I. et al. Incidence of histamine-forming bacteria and histamine content in scombroid fish species from retail markets in the Barcelona area. Int. J. Food Microbiol., 28, 411, 1996. 70. Schubert, R.H.W. Ueber den Nachweis von Plesiomonas shigelloides Habs and Schubert, 1962, und ein Elektivmedium, den Inositol-Brillantgrun-Gallesalz-Agar. Ernst RodenwaldtArch., 4, 97, 1977. 71. Schubert, R.H.W. Genus IV. Plesiomonas. In Bergey’s Manual of Systematic Bacteriology, Vol. 1, Krieg, N.R. and Holt, J.G., Eds. The Williams & Wilkins Co., Baltimore, MD, 1984, 72. von Graevenitz, A. and Bucher, C. Evaluation of differential and selective media for isolation of Aeromonas and Plesiomonas spp. from human feces. J. Clin. Microbiol., 17, 16, 1983. 73. Jeppesen, C. Media for Aeromonas spp., Plesiomonas shigelloides and Pseudomonas spp. from food and environment. Int. J. Food Microbiol., 26, 25, 1995. 74. Miller, M.A. and Koburger, J.A. Evaluation of Inositol Brilliant Green Bile Salts and Plesiomonas Agars for recovery of Plesiomonas shigelloides from aquatic samples in a seasonal survey of the Suwanee river estuary. J. Food Prot., 49, 274, 1985.

415 75. Huq, A. et al. Optimal growth temperature for the isolation of Plesiomonas shigelloides, using various selective and differential agars. Can. J. Microbiol., 37, 800, 1991. 76. Freund, S.M., Koburger, J.A., and Wei, C.I. Enhanced recovery of Plesiomonas shigelloides following an enrichment technique. J. Food Prot., 51, 110, 1988. 77. Varnam, A.H. and Evans, M.G. Foodborne Pathogens: an Illustrated Text. Wolfe, London, 1991. 78. O’Hara, C.M. Evaluation of the Phoenix 100 ID/AST system and NID panel for identification of Enterobacteriaceae, Vibrionaceae, and commonly isolated nonenteric gram-negative bacilli. J. Clin. Microbiol., 44, 928, 2006. 79. González-Rey, C. et al. Specific detection of Plesiomonas shigelloides isolated from aquatic environments, animals and human diarrhoeal cases by PCR based on 23S rRNA gene. FEMS Immunol. Med. Microbiol., 29, 107, 2000. 80. Gu, W.M. and Levin, R.E. Quantitative detection of Plesiomonas shigelloides in clam and oyster tissue by PCR. Int. J. Food Microbiol., 111, 81, 2006. 81. Gu, W. and Levin, R.E. Quantification of viable Plesiomonas shigelloides in a mixture of viable and dead cells using ethidium bromide monoazide and conventional PCR. Food Biotechnology, 21, 145, 2007. 82. Loh, J.P. and Yap, E.P.H. Rapid and specific detection of Plesiomonas shigelloides directly from stool by LightCycler PCR. In Rapid Cycle Real Time PCR. Methods and Applicati­ ons. Microbiology and Food Analysis, Reischl, U., Wittwer, C., and Cockerill, F., Eds. Springer-Verlag, Berlin, 2002. 83. Nair, G.B. and Holmes, B. International Committee on Systematic Bacteriology Subcommittee on the Taxonomy of Vibrionaceae. Minutes of the closed meeting, 19 May 1998, Atlanta, GA, USA. Int J Syst Bacteriol, 49, 1945, 1999. 84. González-Rey, C. et al. Serotypes and anti-microbial susceptibility of Plesiomonas shigelloides isolates from humans, animals and aquatic environments in different countries. Comp. Immunol. Microbiol. Infect. Dis., 27, 129, 2004. 85. Ciznar, I. et al. Potential virulence-associated properties of Plesiomonas shigelloides strains. Folia Microbiol. (Praha), 49, 543, 2004. 86. Gu, W. et al. Genetic variability among isolates of Plesiomonas shigelloides from fish, human clinical sources and fresh water, determined by RAPD typing. Food Biotechnol., 20, 1, 2006.

30 Proteus

Antoni Róz∙alski and Paweł Sta˛ czek University of Łódz´

Contents 30.1 Introduction.....................................................................................................................................................................417 30.1.1 Taxonomy...........................................................................................................................................................417 30.1.2 Habitat and Pathogenicity..................................................................................................................................417 30.1.3 Swarming Phenomenon and Virulence Factors of Proteus Bacteria................................................................418 30.1.4 Conventional Diagnosis.................................................................................................................................... 420 30.1.4.1 Biochemical Tests............................................................................................................................ 420 30.1.4.2 Commercial Systems....................................................................................................................... 420 30.1.4.3 The Dienes Test................................................................................................................................ 420 30.1.4.4 Bacteriophage and Bacteriocin Typing............................................................................................ 420 30.1.4.5 Immunodiagnosis............................................................................................................................. 421 30.1.5 Molecular Diagnosis......................................................................................................................................... 421 30.1.5.1 Amplification of Specific Sequences by PCR.................................................................................. 422 30.1.5.2 Oligonucleotide Arrays.................................................................................................................... 424 30.1.5.3 Proteus Typing................................................................................................................................. 425 30.2 Methods.......................................................................................................................................................................... 426 30.2.1 Reagents and Equipment................................................................................................................................... 426 30.2.2 Sample Preparation........................................................................................................................................... 426 30.2.3 Detection Procedure......................................................................................................................................... 426 30.3 Conclusions and Future Perspectives............................................................................................................................. 427 References.................................................................................................................................................................................. 428

30.1  Introduction 30.1.1 Taxonomy Proteus belongs to the Enterobacteriaceae family. The first description of Proteus rods was made by Hauser in 1885.1 The classification of bacteria in the genus Proteus has been changing throughout the twentieth century. Due to close similarity of Proteus, Morganella, and Providencia rods, all three genera were placed in the tribe Proteeae, however, the tribe designation is not often used.2 The first two species Proteus mirabilis and Proteus vulgaris were originally described by Hauser. In 1966 Cosenza and Podgwaite3 isolated, from larvae of the gypsy moth, bacteria producing slime and biochemically distinct from both P. mirabilis and P. vulgaris. These authors proposed to name these bacteria P. myxofaciens. The biochemical as well as DNA–DNA hybridization analyses showed the homogeneity of P. mirabilis and heterogeneity of the P. vulgaris species. Hickmann et al.4 divided P. vulgaris into three biogroups based on indol production, esculin hydrolysis and fermentation of salicin. The biogroup 1, also defined as genomospecies 1, was indole, salicin, and esculin negative and was named Proteus penneri. Subsequently the additional two P. vulgaris biogrups (2 and 3) were studied. Results of the studies

suggested that biogroup 2 was homogenous and represented a single species ­(genomospecies 2). The name P. vulgaris was reassigned to this species.5 The biogroup 3 consisted of four genomospecies (3, 4, 5, and 6). The studies of O’Hara et al.6 showed that genomospecies 3 contained only two strains exhibiting low relatedness to other P. vulgaris genomospecies and had unique biochemical features (DNase, lipase, and tartrate negative). These results allowed the authors to separate the genomospecies 3 from the other genomospecies as well as to propose that it should be named Proteus hauseri, whereas the other genomospecies remained unnamed as Proteus genomospecies 4, 5, and 6. Thus, the genus Proteus currently consists of five species: P. mirabilis (type strains ATCC 29906), P. vulgaris (ATCC 29905), P. penneri (ATCC 33519), P. hauseri (ATCC 700826 and ATCC 13315) P. myxofaciens (ATCC 19692), as well as Proteus genomospecies 4 (ATCC 51469), 5 (ATCC 51470) and 6 (ATCC 51471).5,7 P. myxofaciens is the only Proteus species without any significance in the pathogenicity of human beings.7

30.1.2 Habitat and Pathogenicity Proteus rods are opportunistic pathogens, which cause different types of infections including urinary tract infections 417

418

(UTI), wounds infections, meningitis in neonates or infants, rheumatoid arthritis (RA) and gastroenteritis, however, these bacteria are rarely associated with foodborne infections.7 Proteus rods are widespread in the natural environment, they can be found in polluted water, soil, and manure. Due to the proteolytic activity, the ability to hydrolyse urea to ammonia and carbon dioxide, as well as the oxidative deamination of amino acids, these bacteria play an important role in decomposing organic matter of animal origin. Proteus bacteria are also a part of the natural gastrointestinal tract flora of humans and animals.2 Therefore, they are frequently isolated from human and animal feces, as well as from the associated sources such as sewage and decomposing meat. P. mirabilis is a common inhabitant of dogs, cows, and birds. P. vulgaris is often recovered from cows and birds, however, in contrast to P. mirabilis, this species is more frequently isolated from pigs than from dogs.8 Human feces were found as a source of nosocomial infections caused by P. mirabilis.9 P. vulgaris and P. penneri were isolated from stools with a wide range of frequencies: 0.45–6% and 0.4–1.2%, respectively.7 There is no strong evidence to classify Proteus rods as enteropathogens. The possible role of Proteus in diarrheic diseases was studied in the 1950s.10 Muller et al.11,12 found P. mirabilis and P. vulgaris in diarrheic specimens and these species were more often isolated from patients with diarrheic diseases than from healthy persons. There was no proof that Proteus strains classified into different serogroups could initiate epidemic enteric diseases. Proteus species were also isolated from food products. Von Holy et al.13 showed that these bacteria can be found in manufactured vacuum packed sausages. Proteus, as well as Serratia and Enterobacter rods, were the three most frequently isolated genera accounting for approximately 4% of total microbial contamination of this product. Proteus rods are commonly isolated from freshwater and saltwater fish.14–17 It was found that these bacteria can produce histamine as a product of histidine decarboxylation during fish spoilage. Contaminated fish consumption leads to the disease known as scombroid poisoning.14,16 Recently P. mirablis rods have been described as the bacteria accumulated in oysters collected from marine coastal area of Venezuela.18 The presence of these bacteria may be attributed to fecal contamination of this area, particularly during rainy periods. Proteus spp. was identified in the samples of raw milk illegally purchased in two major cities in Ghana. Most of identified microorganisms were enterobacteria indicating the fecal contamination of milk due to poor hygiene.19 Recently described studies have shown that P. vulgaris present on the surface of ripe French cheeses had the greatest capacity to produce high quantities and wide varieties of volatile compounds, which play a major role in cheese flavor formation.20 Proteus rods play a particularly important role in UTI. They frequently cause UTI in patients with urinary catheter in place or with structural and/or functional abnormalities within a urinary tract, as well as after surgical intervention in the urogenital system. P. mirabilis causes UTI with the highest frequency among all Proteus species. It is involved

Molecular Detection of Foodborne Pathogens

in complicated infections and infections in long ­catheterized patients. Proteus bacteria can cause two types of UTI— hematogenous infections and ascending infections, however, the latter are more common to these microorganisms. Proteus rods are also associated with nosocomial infections.7,21 Microbiological, molecular, and immunological studies carried out by several research groups support the role of P. mirabilis in the etiopathogenesis of RA. Moreover, the correlation between upper UTI and RA was found due to P. mirabilis.22

30.1.3 Swarming Phenomenon and Virulence Factors of Proteus Bacteria Proteus are dimorphic bacteria, which are able to display two types of behavior. While growing in liquid media they are motile, peritrichously flagellated short rods, 1.0–2.0 µm in length with six to ten flagella. These short bacteria, called swimmer cells, when transferred to a solid medium, differentiate to elongated multinucleated, nonseptated, highly flagellated swarmer cells of 20–80 µm length. The population of swarmer cells can migrate in a highly coordinated manner on the solid media. The migration of the bacterial mass takes place as long as the number of swarmer cells in the population is reduced by loss of individual cells during moving on the surface. Then the long swarmer cells disintegrate into short rods in the process called consolidation. The differentiation of swimmer cells to swarmer cells, their migration on solid media and de-differentiation to short rods, known as a swarming phenomenon or a swarming growth, is cyclical. The repeating cycle of differentiation of short rods and disintegration of swarmer cells result in the formation of characteristic rings that form a bull’s-eye pattern on agar plates23 (Figure 30.1). Swarmer cells differentiation is initiated by the contact of bacteria with the solid surface, by the inhibition of flagellar rotation and it is dependent on multicellular interaction and cell–cell signaling. An extracellular acidic polysaccharide designated colony migration factor (Cmf) which act as lubricant for cells during swarming, facilitates swarmer cells translocation on the solid surface.24 The number of factors involved in the swarming regulation was reviewed by Rather.25 Both morphologically and physiologically different cells—swimmer short rods and swarmer elongated cells—play an important role in pathogenesis. Highly flagellated swarmer cells are thought to be crucial in ascending UTI infections, whereas short rods containing fimbriae are responsible for colonization of host mucosal surface.26 Proteus has evolved multiple virulence factors including fimbriae, flagella, enzymes (urease, proteases, deaminases), toxins—hemolysins and endotoxin (lipopolysaccharide, LPS) and invasiveness.27–30 These bacteria posses six types of fimbriae or afimbrial adhesins: mannose resistant Proteus like fimbriae (MR/P), mannose resistant Klebsiella hemagglutinins (MR/K), Proteus mirabilis fimbriae (PMF), nonagglutinating fimbriae (NAF), ambient temperature fimbriae (ATF) and P. mirabilis P-like fimbriae (fimbriae, which are similar

Proteus

Figure 30.1  Swarming of P. mirabilis on nutrient broth agar plate as a form of concentric rings of growth. (From Kwil, I., et al., unpublished data, 2006.)

to E. coli P pili).27 MR/P fimbriae facilitate colonization of upper urinary tract and are more often found on strains which cause pyelonephritis.28 MR/K hemagglutinins are associated with adhesion of bacteria to catheters. PMF are involved in colonization of both a bladder and kidneys. NAF was previously described as uroepithelial cell adhesin (UCA). ATF as well as P. mirabilis P-like fimbriae are not important in the pathogenicity.29,30 Flagella are important bacterial surface structures which facilitate microbial colonization of the infected host. They play a crucial role in swarming phenomenon of Proteus.25 Flagellin, which forms flagellar filament is a strongly immunogenic H-antigen, inducing immunoresponse of the macroorganism. Bacteria possessing the mechanism of flagellar variation can escape the immunoresponse. Indeed, due to a rearrangement of genes encoding flagellin P. mirabilis can produce six classes of antigenically and biochemically different flagellin, which distribution on short rods and swarmer cells is different. This kind of antigenic variation most probably protect bacteria against the activity of secreted class A immunoglobulins (sIgA), which are produced by uroepithelial cells to inhibit their dissemination by immunobillization.31 Proteus can also escape the action of sIgA by production of cell free proteases, which degrade both types of IgA and also IgG antibodies, as well as other types of proteins including complement.32,33 Two Proteus proteases ZapA and ZapD were described.34 Mutants, which cannot produce these enzymes, turned out to be less virulent compared to wild type strains.35 Most probably proteases in vivo diminish phagocytosis due

419

to the hydrolysis of opsonic antibodies to fragments unable to bind to phagocytic cells by Fc fragment.36 Urease, an enzyme digesting urea to ammonia and carbon dioxide, was demonstrated as the most important virulence factor of Proteus strains.27–30 It was found that urease negative mutants were less virulent than the parental strains.37 The urease action during UTI leads to the pH elevation which results in the precipitation of magnesium and calcium ions and to the formation of urinary stones containing struvite and carbonate apatite.29 The acidic bacterial polysaccharides (capsule, O-antigens) may be also involved in urinary stones formation.38,39 The third type of enzymes important for Proteus infections are amino acids deaminases producing α-keto acids which play the role of siderophores for these bacteria.40 Proteus rods synthesise cell-associated calcium independent HpmA hemolysin which is a 166-kDa protein.41 This toxin seems to be not so important in pathogenesis of this genus which was shown by use of HpmA– mutant.28 Three species P. vulgaris,42 P. penneri,43 and P. hauseri 44 also produce cell free, calcium dependent HlyA hemolysin classified into RTX proteins (repeated in toxin). Both hemolysins belong to the family of pore forming toxins, which are able to act not only against erythrocytes but also can destroy other types of cells.29 Cell invasiveness, a characteristic feature of Proteus bacteria, indicated the possible role of these microorganisms as enteropathogens. Several authors have shown the ability of P. mirabilis, P. vulgaris, and P. penneri to penetrate different cell lines including Vero (the African green monkey kidneys cells), HeLa (the cancer cells of the uterus neck), L929 mouse fibroblast, human bladder cell lines (T24, EJ/28 and 5637), Hu 609 (human ureteric ueroepithelial cell lines), HRPTEC (human renal proximal tubular epithelial cells), Int 407 (embryonic intestinal epithelial cells) and HCT-8 (ileocecal epithelial cells).7 It was found that P. mirabilis strains—highly active in production of HpmA hemolysin— are the most invasive.45,46 The penetration of cell lines by P. vulgaris and P. penneri strains producing HlyA hemolysin is accompanied by the cytotoxic effect of this cytolysin.43 The mechanism of Proteus invasiveness has not been elucidated. It was demonstrated that internalized bacteria in HRPTEC were present within membrane bound vacuoles and that microfilament formation is not involved in the internalization process due to the actin polymerization.47 Some authors found that the bacteria can replicate within the cell lines,47 whereas the others did not confirm this.46,48 Conflicting results were also obtained while testing the invasiveness capacity of swarmer cells. Allison et al.48 showed that the invasiveness ability of Proteus is closely related to the swarming phenomenon and the swarmer cells are more efficient invaders than short rods. Other authors46,47 found that short vegetative cells which number increased due to the reversion of swarmer long bacteria in liquid medium were internalized. Proteus, as with all Gram-negative bacteria, contain LPS in the outer membrane which when released from bacteria is

420

a biologically active endotoxin. LPS causes a broad spectrum of pathophysiological effects, particularly septic shock.49 Proteus LPS contains three parts: O-specific polysaccharide (O-antigen), core region and lipid A, the last one is an ­endotoxin biological center. The differences in the structure of O-antigens serve as the basis for the serological classification of Proteus strains50 (see below).

30.1.4 Conventional Diagnosis 30.1.4.1 Biochemical Tests Proteus spp. is easily recognized on media due to their ability to swarm on agar media surfaces. Microscopically swarming phenomenon appears as concentric rings of growth starting from the colony or inoculum site7,23,51 (Figure 30.1). Since almost all Proteus strains typically swarm, they are visible on most of common agar media used in Enterobacteriaceae diagnosis, such as MacConkey, Hektoen enteric, S–S (Salmonella–Shigella) and xylose-lysine-deoxycholate.7,52 Swarming phenomenon makes it difficult to isolate non-Proteus bacteria growing on the same media. Therefore, a number of substances are suggested to inhibit swarming, including phenol, sodium azide, sodium salicylate, ρ-nitrophenyl glycerol, bile salts and high concentration of agar.7 A number of selective media were specifically developed for Proteeae isolation. Hawkey et al.53 described the medium called PIM (Proteeae isolation medium). It is a selective and differential medium containing clindamycin and colistin as selective agents and tryptophan/tyrosine as differential ones. PIM is useful to differentiate Proteeae from contaminated bacteria in clinical specimens. Bacteria of this tribe degrade tyrosine and produce a reddish-brown pigment containing melanin due to the action of enzyme tyrosine phenol-lyase. The selective medium called selective Proteeae medium (SPM) was described by Urbanova.54 Identification of Proteeae on SPM is based on the detection of the phosphatase activity, which is produced by bacteria. This medium contains a tryptose phosphate with phenylphtalein monophosphate as the substrate, bile salts and polymyxin B as inhibitors of enteric bacteria and methyl green as indicator. P. mirabilis and P. vulgaris grow on SPM as light green colonies. Bacteria from the tribe Proteeae can be differentiated by biochemical tests listed in Table 30.1. Members of the Proteus genus are lactose negative, urease, and phenylalanine deaminase positive. They also produce hydrogen sulfide and oxidatively deaminated tryptophan.7,51,52 As it is shown in Table 30.2, six tests—indole production, ornithine decarboxylase, maltose, salicin, and xylose fermentation, esculin hydrolysis—are used to identify the Proteus species isolated from clinical materials and for identification of nonpathogenic P. myxofaciens.55 It is necessary to underline that both P. mirabilis and P. penneri are indole negative, however, P. penneri and 25% of P. mirabilis strains produce a green color when tested by Kovacs’ test.56 Susceptibility to chloramphenicol is also suggested to be useful in identification of the Proteus species.57

Molecular Detection of Foodborne Pathogens

Table 30.1 Differentiation of the Three Genera in the Proteeae Proteus

Providencia

Morganella

Citrate utilization

v

+

-

D-Mannose fermentation

-

+

H2S production

-

+ v

Ornithine decarboxylese

+ v

Urea hydrolysis Inositol fermentation

Test

(+)

+

V

-

V

-

Gelatin liquefactions (22°C) Lipase (corn oil)

+

-

-

+

-

-

Swarming

+

-

-

+

Source: Janda, J.M. and Abbot S.L., In The Enterobacteriaceae, 2nd ed., ASM Press, Washington, D.C., 2006; Hawkey, P.M., In Principles and Practice in Clinical Bacteriology, 2nd ed., Gillespie, S.H. and Hawkey P.M., Eds. John Wiley & Sons, Ltd. England, 2006; Weissfeld, A.S. et al., In Clinical and Pathogenic Microbiology, 2nd ed., Howard, B.J. et al., Eds., Mosby, st Louis, 1994; Senior, B.W., J. Med. Microbiol., 46, 39, 1997. Notes: +, 90–100% positive; (+), 75–89.9% positive; v, 25.1–74.9% positive; -, 10.1–25% positive.

30.1.4.2  Commercial Systems Several rapid, automated, and manual systems of identification of Proteus bacteria at species level are available. It was reported that most of systems, including Crystal ID-E/NF (Becton Dickinson Microbiology Systems), GNI, GNI+ and API 20E (BioMerieux Inc.), Rapid Neg ID3 (Dade Behring Inc.), Vitek GNI card and Vitek GNI+ card (BioMerieux Vitek) are highly accurate (>90%) in identification of three clinically significant species: P. mirabilis, P. vulgaris and P. penneri, however the latter may cause identification errors. Misidentification within Proteus sp. as well as identification of Proteus bacteria as Morganella morganii or Providencia rettgeri have been reported.5,7 30.1.4.3 The Dienes Test One of the oldest assay applied to epidemiological typing is the Dienes test.7,21,53 When two different strains of Proteus spp. are inoculated onto an agar plate and allowed to swarm toward each other, a demarcation line of growth appears if the strains are incompatible. Such reaction does not occur if the strains are identical. The exact basis for the Dienes phenomenon is not established, however, it is believed that it is related to production and susceptibility to bacteriocins (proticins) by Proteus strains. The Dienes test is a method easily applied for epidemiological investigations and yields highly discriminatory results.21 30.1.4.4 Bacteriophage and Bacteriocin Typing Bacteriophage and bacteriocin typing are not routinely used in clinical laboratory work. There are three schemes of phage typing described for Proteus and other members of Proteeae, P. mirabilis, and P. vulgaris, as well as only for

421

Proteus

Table 30.2 Differentiation within the Genus Proteus P. mirabilis

P. vulgaris

P. penneri

P. hauseri

P. myxofaciens

Indole production

-

+

-

+

-

Ornithine decarboxylase

+

-

-

-

-

Maltose fermentation

-

+

+

+

+

Salicin fermentation

-

+

-

-

-

D-Xylose fermentation

+ –

+

+

+

-

Esculin hydrolysis Chloramphenicol susceptibility

S

+ V

R

S

S

Test/Property

Source: Janda, J.M. and Abbot S.L., In The Enterobacteriaceae, 2nd ed., ASM Press, Washington, D.C., 2006; Hawkey, P.M., In Principles and Practice in Clinical Bacteriology, 2nd ed., Gillespie, S.H. and Hawkey P.M., Eds. John Wiley & Sons, Ltd. England, 2006; Weissfeld, A.S. et al., In Clinical and Pathogenic Microbiology, 2nd ed., Howard, B.J. et al., Eds., Mosby, st Louis, 1994; Senior, B.W., J. Med. Microbiol., 46, 39, 1997. Notes: +, 90–100% positive; -, 0–9.9% positive; S, susceptible; R, resistant; V, variable susceptibility.

P. mirabilis strains.21 Hickman and Farmer58 differentiated 200 P. mirabilis strains into 113 types using 23 lytic bacteriophages. Senior59 examined Proteus strains for production of proticins (P) as well as proticins sensitivity (S) and found that by use of this assay 250 strains can de differentiated into 90 P/S types. The P/S type of Proteus isolate is independent of O serogrouping.60 30.1.4.5  Immunodiagnosis Proteus sp. is antigenically heterogenous, particularly due to structural differences in the O-specific part (O-antigen) of LPS, as well as in H-antigen (flagella). The differences in the O-antigen structure serve as the basis for the serological classification of Proteus bacteria. The Kaufman and Perch scheme includes 49 different P. mirabilis and P. vulgaris O-serogroups and 19 serologically distinct H-antigens.61 This classification was then supplemented by Larson,62 as well as by Penner and Hennesy63 with additional serogroups. The chemical and serological studies conducted subsequently allowed additional O-serogroups to be established, which classified P. penneri and P. hauseri strains as well as P. myxofaciens and some P. mirabilis and P. vulgaris strains unclassified by the above mentioned schemes.64,65 The serological classification currently consists of 76 serogroup.50,66 The most frequently isolated Proteus strains in clinical infections were classified into serogroups O3, O6, O10, O26, O27, O28, and O30.21,62 The serological specificity of Proteus O-antigens was studied using the polyclonal rabbit anti-O sera specific to the particular serogroups.29,50,65,66 Polyclonal anti-O sera contains antibodies of different specificity types. Usually, the major antibodies fraction recognizes the main epitope, which defines the group specificity, whereas the minor fractions can bind other epitopes in O-antigen or in core region of LPS. The agglutination test for the determination of O-antigens should be performed with cultures that were boiled for 1 h to destroy the H-antigen.21 The study of the antisera specificity can also be conducted using the isolated LPS representing particular serogroups and enzyme immunosorbent assay (EIA), as well as passive hemolysis test (PHT).67–69

Uronic acids and hexosamines, the characteristic compounds of Proteus OPSs, play an important role in the serological specificity of these antigens.29,30,50 α-D-GlcA and β-D-GlcA present as a branch of O-specific polysaccharides chain, as well as in linear O-polysaccharides usually serve as immunodominant sugars in specific epitopes.67,70,71 The most common epitopes showed for Proteus O-antigens were uronic acids substituted by amino acids. The importance of α-D-GalA(-L-Lys) in the specificity of P. mirabilis O3,72 O26,73 and O28,74 as well as α-D-GalA(-L-Thr) in the specificity of P. mirabilis O1175 were described. O-specific antibodies may cross-react with LPS of strains belonging to the same species or genus but classified into other serogroups. Such cross-reactivity occurs also even with the LPSs of taxonomically different bacteria. For example, due to the common α-L-FucNAc-(1→3)-α/β-D-GlcNAc disaccharide a cross reactivity was observed in antigen–­antibodies systems of LPS and heterologous antisera of P. vulgaris O8, O12, O39, and P. mirabilis O6.50 Proteus O-antigens also show similarity with OPSs from bacteria of other genera. For example, cross-reactivity was found between the anti-P. mirabilis O13 serum, as well as the anti-P. myxofaciens serum and the LPS from Providencia alcalifaciens O14 and O23. Serological studies revealed an important role of common compound alanino-lysine (Nε -[(R)-1-carboxyethyl]L-lysine (alanino-lysine), AlaLys) for the specificity of these O-antigens.76,77 The common compound D-(4-amino4,6-dideoxyglucose (4-aminoquinovose), Qui4N) carrying N-linked N-acetyl-aspartic acid is responsible for the antigenic relationship of O-antigens of P. mirabilis O38, as well as P. alcalifaciens O4 and O33.78,79 The knowledge of the chemical Proteus O-antigens structure as well as the specificity of O-antisera enables mistakes to be avoided in immunodiagnosis of these bacteria.

30.1.5 Molecular Diagnosis Although many molecular detection and typing techniques of that group of bacteria have been developed, specific,

422

standardized procedures for analysis of Proteus in clinical specimens and food have not been established. Among many obstacles found during the application of molecular techniques to pathogen detection (e.g., a problem of amplification of the DNA originating from the dead cells), the presence of accompanying bacterial flora plays an important role. The number of pathogens of interest may be significantly lower than the number of indigenous bacteria, thus the applied tests should be highly sensitive and specific. Most of the methods described below and summarized in Table 30.3 have been tested on the bacterial cultures raised from single colonies of the pathogen obtained during preliminary procedures based on the classical microbiologic strategies or on clinical samples of relatively low microbial complexity. Nevertheless, their efficiency, reliability, and universal character allow one to presume that these methods will find their place in the guideline schemes for analysis of contaminated samples. 30.1.5.1 Amplification of Specific Sequences by PCR This group of tests utilizes specific primers allowing for the amplification of group- or species-specific regions. The well-characterized 16S rRNA gene is the most common target region utilized in pathogen detection. It contains variable regions flanked by highly conservative sequences which are very useful in primer design. Using this approach P. mirabilis has been detected as a predominant pathogen colonizing two species of oysters collected from the vacation area at the Venezuelan coast.18 Phosphate-buffered saline (PBS) homogenized oyster tissues were used to inoculate suitable microbiological media (McConkey, Nutrient Broth as well as selective Chocolate Agar) and selected liquid cultures were used for DNA isolation followed by amplification of 16S rDNA using primers harbored in the highly conservative regions. The amplified fragments of ~1.4 kb were subsequently sequenced, which identified the predominant species to be highly similar (99%) to P. mirabilis. This method, although very precise in identification of the strains, is very sensitive to any DNA contamination, thus it requires pure bacterial cultures and may be used only as the confirmation to the standard microbiological tests. PCR fragments of the amplified 16S rDNA may be also analyzed using restriction fragment length polymorphism. Lu et al.80 used this method to differentiate common pathogens isolated from body fluids. Fragments obtained with a pair of universal primers were digested with HaeIII endonuclease, yielding specific pattern for three P. mirabilis strains, distinguishing this species from the other 17 pathogens tested. The sensitivity of the assay was estimated for >10 CFU/1 ml in the case of Gram-negative bacteria. Identification of the pathogen in life-threatening cases, such as septicemia, requires development of methods which are not only reliable, but quickly provide results. Vliegen et al.81 combined amplification of the 16S rDNA with realtime PCR methodology which allows for the monitoring of a specific product formation during amplification. Using this technology 94 different bacterial species were tested,

Molecular Detection of Foodborne Pathogens

including P. mirabilis and P. vulgaris strains, and the whole ­identification procedure lasted 6.5–7 h. Proteus bacilli were also identified using analysis of ­single-stranded DNA conformation polymorphism (SSCP) of the amplified 16S rDNA regions. Amplicons were subjected to heat denaturation in order to obtain single strands which were subsequently resolved electrophoretically in the nondenaturing polyacrylamide gel. Electrophoretic mobility of the fragments is a consequence of the specific nucleotide sequence of the variable regions which determines the unique spatial conformation of the separated strands. Multiplex PCR utilizing a set of primers targeted to the conservative regions allowed, in case of Proteus, to detect specific patterns distinguishing P. vulgaris and P. mirabilis from 47 different pathogen species.82 Sensitivity, repeatability, and resolution were improved by fluorescence labeling of the primers, which allows for the polymorphism detection using automatic DNA sequencer. Each lane is thus analyzed at the bottom by a sensor, which detects laser-induced fluorescence signals, and the differences in the mobility of particular single-stranded DNA molecules are expressed as the shift in the retention time so the data may be simply processed and stored in the computer. Moreover, the use of an automatic sequencer allows strict control of gel temperature as well as the addition of internal markers which significantly improves the repeatability of the results. Sensitivity of this method was estimated at >101 CFU of bacteria in the samples consisting pure cultures and the whole procedure lasted only 8 h (excluding cultivation and DNA isolation). 16S rDNA was also used as a target by SchebereiterGurtner et al.83 to identify pathogens responsible for conjunctivitis. In this case, amplification was followed by denaturing gradient gel electrophoresis (DGGE). During DGGE short amplicons of 16S rDNA variable regions migrate toward increasing concentration of denaturing agent, leading to the partial breakage of hydrogen bonds which results in the reduced mobility of the molecules. As in the SSCP technique, specific patterns are obtained which may be used for the identification as well as genotyping of bacteria. Among 60 samples collected from human conjunctivae infected with monomicrobial and polymicrobial communities, 27 pathogenic strains were detected including one identified as Proteus sp. Consequent sequencing of the PCR product affiliated this strain to P. mirabilis. Along with the techniques utilizing 16S rDNA for the identification of bacterial species, methods focused on the different other target regions are being developed. Mansy et al.84 designed primers based on the sequence of the previously cloned P. mirabilis specific DNA fragment. Using these primers, chromosomal DNAs of 99 strains belonging to almost 40 different species were analyzed, revealing the presence of a specific 3.5 kb band only for the members of P. mirabilis species. The minimal sensitivity of the test was estimated approximately for 10 fg of DNA which corresponds to ~17 bacterial cells, although only pure cultures were tested in this analysis. PCR amplification of urease ureC1 and ureC2 elements specific for P. mirabilis was used for detection of this

Microsatellite tetramer

Repetitive DNA

P. mirabilis specific DNA fragment

23S rDNA

16S rDNA, 23S rDNA

MMKAP 1 ACCTAGTCCCAGAAAAGAACCCTC MMKAP 2 TGGTGTTAATCACATGCTTGATGG Rep1R-Dt IIINCGNCGNCATCNGGC Rep2-Dt NCGNCTTATCNGGCCCAT BoxA1R CTACGGCAAGGCGACGCTGACG GACAGACAGACAGACA or CAATCAATCAATCAAT

Capture probe GGGTTCATCCGATCGTGCAAGGTCCGAAGAGCCCC Detector probe GGTCCGTAGACATTATGCGGTATTAGCCACCGTTT 16S-F1 CGCTGGCGGCAGGCCTAACACATGC 16S-R1 Cy3-CGCGGCTGCTGGCACGGAGTTAGCC 23S-F3 ACCGATAGTGAACCAGTACCGTGAG 23S-R4 Cy3-TTAAATGATGGCTGCTTCTAAGCC Proteus spp. probe CGAGCGGTAACAGGAGAAAGCTTGCTTGCTTTCTTGCTGA P1 ACCAGGATTTTGGCTTAGAAG P2 CACTTACCCCGACAAGGAAT P. vulgaris probe GTAGGCAGAGTGATTAGGCAAA

16S rDNA

16S rDNA

16S rRNA

Primer and Probe Sequences (5′–3′)

Amplification primers 8Fpl AGAGTTTGATCCCTCAG 1525R AAGGAGGTGATCCAGCC Internal sequencing primers 515F GTGCCAGCMGCCGCGGTAA 519R GWATTACCGCGGCKGCTG 926F GGGCCCGCACAAGCGGT 926R ACCGCTTGTGCGGGCCC P11P GAGGAAGGTGGGGATGACGT P13P AGGCCCGGGAACGTATTCAC ER10 GGCGGACGGGTGAGTAA ER11G GGACTGCTGCCTCCCGTAG ER14 GCTAACTCCGTGCCAGCA ER15 GCGTGGACTACCAGGGTATC U1 CCAGCAGCCGCGGTAATACG U2 ATCGGC/TTACCTTGTTACGACTTC

16S rDNA

Target Sequence

Table 30.3 Selected Protocols used in Molecular Identification and Typing of Proteus Strains

1×95°C/7’ 32×(92°C/30’’, 40°C/1’, 65°C/2’) 1×65°C/16’

1×94°C/7’ 32×(94°C/30’’, 40°C/1’, 65°C/2’15’’)

1×94°C/5’ 35×(94°C/1’, 58°C/1’, 72°C/2’) 1×72°C/7’

1×95°C/5’ 5×(95°C/40’’, 48°C/40’’, 72°C/50’’) 25×(95°C/40’’, 53°C/45’’, 72°C/50’’) 1×72°C/10’

1×37°C/10’ 1×94°C/5’ 35×(94°C/30’’, 56°C/30’’, 72°C/30’’) 1×72°C/5’

1×94°C/10’ 35×(94°C/1’, 55°C/1’, 72°C/2’) 1×72°C/10’ None

25×(94°C/1’, 55°C/1’, 72°C/10’’)

1×94°C/6’ 30×(94°C/45’’, 55°C/45’’, 72°C/1’) 1×72°C/10’

Cycling Program

P. mirabilis typing

Proteus identification and typing

P. mirabilis detection by amplification of species-specific region

P. vulgaris detection using microarrays

Proteus sp. detection using microarrays

Detection of P. mirabilis using microarrays

Detection of P. mirabilis by PCR-RFLP (HaeIII endonuclease)

107

93

84

94

95

96

80

82

Detection of Proteus by multiplex PCR followed by SSCP analysis

Reference 18

Strategy Detection of Proteus by 16S rDNA amplification and sequencing

Proteus 423

424

microorganism in urinary calculi. 87 stones were used for DNA extraction by boiling pulverized pieces, revealing the presence of P. mirabilis in struvite stones, but not in metabolic ones (calcium oxalate, calcium phosphate, uric acid, cystine) and giving a higher detection rate than the stone and urine culturing method.85 Kwil et al.86 reported a detection strategy in which the primers were based on the internal sequence of hpmA gene from Proteus mirabilis, but the amplification of the chromosomal DNA was carried out under conditions allowing for the formation of nonspecific random products. Characteristic common patterns of randomly amplified polymorphic DNAs (RAPD) were obtained for 69 P. penneri strains which allowed for distinguishing this species from other Proteus strains tested (32 P. vulgaris and 19 P. mirabilis) and seven other pathogens represented by one to 12 strains. The other targets which may be useful for the detection of Proteus species are the sequences encoding class II restriction endonucleases. These enzymes are widely distributed among different groups of bacteria and often show high specificity not only within bacterial family or species but sometimes also differ between serotypes.87–89 Currently over 5000 of such enzymes have been discovered, and the DNA sequence of above 200 genes encoding these enzymes has already been revealed (http://rebase.neb.com). The comparative study of these sequences showed significant differences between them, even if they encoded enzymes very similar in the organization and properties.90 In the case of Proteus genus, two class II restrictases (PvuI and PvuII) genes have been sequenced, both of them specific for P. vulgaris strains. Specific primers derived from the pvuII sequence as well as the other primers targeting species-specific endonucleases have been used for the detection of eight different pathogens among over 100 strains tested by individual and multiplex PCR amplification of chromosomal DNA samples isolated from pure cultures.91 This method although promising for the detection of some pathogens, showed to be too discriminatory in the case of the analyzed 12 P. vulgaris strains as only one strain gave a positive signal. Such results suggest that this enzyme is not widely distributed among Proteus genus, or there are some significant clonal differences within the sequences of pvuII gene. Another PCR based technique which may be utilized for the detection of Proteus is rep-PCR which uses primer oligonucleotides anchored to the repetitive sequences (rep) which are widely distributed in Enterobacteriaceae genomes.92 Amplified products of different sizes are formed due to the genus-, species-, or even clone-specific distribution and orientation of rep sequences, which allows for the identification and genotyping of bacteria. Application of RepDt pair of primers gave distinct amplification patterns for chromosomal DNAs isolated from pure Proteus cultures, among which the bands specific for particular species were detected.93 In the case of P. vulgaris two specific bands, 1880, and 260 bp were distinguishable, whereas for P. mirabilis 560 bp specific band was observed. The most complex set of bands was found for

Molecular Detection of Foodborne Pathogens

P. penneri where seven specific fragments ranging from 240 to 3650 bp were identified. The presence of species ­specific bands was also reported when other rep sequences were used for primer designing, such as ERIC (three bands specific for P.vulgaris—330, 830, 1690 bp; one band for P. mirabilis—740 bp; three bands for P. penneri—700, 960, 1350 bp) BoxA2 (610 bp band for P. vulgaris, 1050 bp for P. mirabilis) and BoxA1 (one specific band for P. vulgaris, P. mirabilis and P. penneri—610, 530, and 2490 bp, respectively).93 30.1.5.2 Oligonucleotide Arrays Although originally developed for gene expression analysis, microarrays gain significant attention as a tool for identification of microorganisms in food, clinical, and environmental samples. Briefly, it is based on the set of specific oligonucleotides fixed to the solid support, which are able to hybridize with the complementary regions of the target sequences amplified previously by PCR or multiplex PCR. The most frequently used targets are variable regions of 16S rDNA, 23S rDNA, 16S/23S rDNA intergenic region, but other regions of conservative genes, prone to accumulate mutations are also good candidates. Specificity of reaction is mainly due to the length of an oligo used as a probe and the degree of conservation of the amplified target. In Proteus diagnostics several types of microarrays have been developed to date, differing in target and probe sequences as well as the strategy of hybridization signal generation and the level of automation in signal detection. In order to avoid advanced technology which would limit the application of the method to the advanced laboratories, Hong et al.94 developed a very simple microarray in which 23S rDNA was used as a target. This region shows relatively high levels of mutation within long variable region, which makes it a useful candidate in microbe detection. A positively charged nylon membrane was divided into 3 × 3 mm regions and species- or genus-specific synthetic oligonucleotides were spotted for particular positions of the grid. 900 bp fragments of 16 pathogens’ 16S rDNAs were amplified using digoxygenin labeled primers and hybridized to the probes. Signals were detected using anti-digoxygenin antibodies coupled with alkaline phosphatase which catalyzed formation of the color product from the nitro blue tetrazolium chloride/5­bromo-4-chloro-3-indolyl-phosphate (NBT/BCIP) substrate. P. vulgaris specific probe allowed for detection of these rods in pure and mixed cultures as well as the real samples from foodborne infections. Sensitivity of reaction was estimated in foodborne infection mock sample which in case of Proteus yielded 101 CFU when the specimen was additionally contaminated with Bacillus cereus and Vibrio cholerae. In order to increase the spectrum of pathogens detected in one procedure it is possible to apply microarray strategy developed by Jin et al.95 where two targets, 16S, and 23S rDNA, amplified in asymmetric multiplex PCR with fluorescein labeled primers, are utilized. Hybridization with 22 probes specific for 15 pathogens species including

Proteus

Proteus spp. was found to be useful for correct detection of all diagnosed bacteria in less then 3 h. Detailed analysis of the reaction sensitivity revealed, that in the case of pure cultures it was possible to detect 103 CFU/1 ml, whereas for the samples consisted of six cultures this value reached 105 CFU/1 ml. More complex strategy was used for the detection of uropathogens in urine samples, in which 16S rRNA was used as a target instead of 16S rDNA.96 Amplification step was thus omitted in this case, as the target is simply released during cell lysis, and hybridized directly with the specific detector probes—fluorescein labeled oligonucleotides. Such complexes were then spotted onto a microchip containing 16 sensors, each consisted of three electrodes (working, reference, and auxillary). Sensor arrays were made of 50 nm layer of gold deposited on the plastic support and isolated by the alkanethiolate self-assembled monolayer (SAM). Activated C-terminus of SAM was biotinylated which allowed for the attachment of streptavidine which was then used for anchoring of biotinylated capture probes. Capture probes, similarly to detector probes, consisted of 27–35 bp long oligonucleotides, showing high group- or speciesspecificity. After deposition of the detector probe-rRNA complexes on each of the sensor, a consecutive hybridization event took place, this time between a specific capture probe and its corresponding region of a complexed rRNA. These ternary complexes, attached to the solid support, were then detected using horseradish peroxidase coupled antibodies raised against fluorescein. After removal of nonspecifically bound antibodies, microarrays were placed in a chip reader in the presence of 3,3′5,5′-tetramethybenzidine (TMB)-H2O2 solution covering the area of each sensor. Application of the fixed potential between the working and reference electrodes led to the electron transfer in the peroxidase-catalyzed red-ox reaction, which could be analyzed amperometrically. The result was thus a function of the number of peroxidase molecules captured indirectly at the sensor surface. Using this method P. mirabilis as well as other uropathogens was detected within less then 1 h, yielding sensitivity of approximately 4 × 10 4 CFU/1 ml of urine, which corresponded to ~3 × 10 –16 rRNA per analysed sample volume. Microarray strategy for which gyrB gene was chosen as a target have been developed and validated by Kostic et al.97 using a set of 20 pathogens, including P. mirabilis. Approximately 1200 bp fragments of this gene were amplified from bacteria and used for specific fluorescence end-labeling of the designed reverse complement (RC) oligonucleotides lacking the 3′ terminal cytosine residue. Only interaction of the RC oligo with the corresponding gyrB fragment allowed for the incorporation of a terminal labeled ddCTP. This way, after subsequent hybridization of the RC oligos to the microarray containing complementary, species- or genus-specific capture oligonucleotides, fluorescent signals were detected. Such an approach reduces the amount of labeled nucleic acids and decreases the level

425

of nonspecific signals, thus improves the sensitivity of the assay, especially when the relative abundance of the target pathogen is low within the microbial community of the analyzed specimen. 30.1.5.3  Proteus Typing Methods of molecular typing of Proteus bacilli have not been applied extensively in practice due to the fact that the Dienes test, described above, is an extremely simple, low cost and easy to interpret assay. In contrast, ribotyping—a commonly used technique for interspecies characterization of many different pathogens, e.g., Legionella,98 Haemophilus influenzae,99 Vibrio cholerae,100 Clostridium,101 Yersinia pseudotuberculosis,102 as well as Proteeae group,103 requires application of specialistic hardware and trained personnel. Moreover, Pfaller et al.104 compared the classical Dienes test with the automated ribotyping and pulse field gel electrophoresis (PFGE) of the SfiI endonuclease digested DNAs, using collection of 63 P. mirabilis clinical isolates. Both ribotyping and Dienes test identified 40 independent clones and the remaining 23 strains were divided into 13 Dienes types, 12 ribotypes and 14 PFGE types. All three methods revealed only three clusters, each containing two strains of identical patterns, whereas other strains showed distinct patterns, thus reaching almost identical discriminatory power. On the other hand, Sabbuba et al.105 utilized PFGE method in which the NotI digested DNAs from 55 cathether-associated P. mirabilis strains were used, indicating higher discriminatory power than the Dienes test. Nevertheless, several other fingerprinting techniques for P. mirabilis, P. vulgaris, as well as P. penneri species have been described. Bingen et al.106 investigated P. mirabilis cross-infection in hospital environments by application of both ribotyping and arbitrarily primed PCR (AP-PCR), finding AP-PCR easier to perform. Application of a single short, ten nucleotide long primer in AP-PCR allowed for the deep characterization of the isolates from 18 pregnant patients and excluded the common source of infection, despite such suggestions from the prior phenotypic analysis. Fingerprinting method based on the microsatellite sequences was proposed by Cieslikowski et al.107 for typing P. mirabilis strains. Two microsatellite sequences (GACA)4 and (CAAT)4 served independently as primers for differentiation of 40 laboratory strains and 42 clinical isolates. Both, (GACA)4 and (CAAT)4 primers yielded high discriminatory power (0.992, and 0.940, respectively) which places them within the range of Dienes test (0.98), ribotyping (0.92), and PFGE (0.979) established by Pfaller et al.104 Several other primers targeting different sequences have been applied to differentiate Proteus species, among them, those based on the repetitive sequences described above like ERIC, BOXA1, BOXA2, REP,93,108 Recently Drzewiecka et al.109 used BoxA1 primer and a pair of Rep-Dt primers to confirm a case of transmission between gastrointestinal and urinary tract of the patient by P. mirabilis strain.

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Molecular Detection of Foodborne Pathogens

30.2  Methods 30.2.1 Reagents and Equipment Reagents Chromosomal DNA Isolation TE buffer (10 mM Tris-Cl, 1 mM EDTA, pH 8), lysozyme, STEP buffer (50 μl Tris-Cl pH 8, 50 μl 10% SDS, 160 μl 0.5 M EDTA pH 8, 1 mg proteinase K, H2O add 1 ml), phenol, chloroform, isoamyl alcohol, sodium acetate, ethanol, RNaseA DNA Amplification Taq DNA polymerase, polymerase buffer, dNTPs, primers, PCR product purification kit Agarose Gel Electrophoresis Agarose, TAE buffer (40 mM Tris-acetate, 1 mM EDTA), ethidium bromide (0.5% solution), loading buffer (0.25% bromophenol blue, 40% sucrose in H2O) RFLP Analysis Restriction endonuclease, endonuclease buffer, ethanol, acrylamide:N,N′-methylenebisacrylamide solution (29:1), TBE buffer (45 mM Tris-borate, 1 mM EDTA)

Equipment and Supplies Incubator Thermal block or water bath Microcentrifuge UV-Vis spectrophotometer Thermocycler Power supply up to 300 V and 200 mA A system for horizontal agarose gel electrophoresis UV-transiluminator A system for vertical polyacrylamide gel electrophoresis Culture tubes 12 ml Eppendorf 1.5 ml tubes Thin-wall 0.2 ml PCR tubes Automatic pipettes: 200–1000 μl, 20–200 μl, 0.5–10 μl Tips: 1000 μl, 200 μl, 10 μl Disposable gloves

30.2.2 Sample Preparation Development of molecular methods for microorganism detection which allow for fast and precise identification of pathogens, paradoxically created a dilemma for microbiologists. At the current stage, though many of these methods reveal very high specificity and the sensitivity at the level of single cells in case of pure bacterial cultures, when it comes to the analysis directly in biological material, usually in the presence of accompanying bacterial flora, these parameters decrease down dramatically, even by several orders. Therefore, evaluation should be made whether the sensitivity range of the proposed detection technique is appropriate to the estimated amount of bacteria and the complexity of their population in the analyzed material, preliminary stages of sample concentration should be considered, and in case of any doubt, full microbiological diagnostics should be performed. In such a case, molecular methods should serve as a reliable confirmation of classical techniques or they should decide in a hard-to-diagnose case. As a method of choice, individually these techniques should be used only in a life threatening cases, e.g., sepsis, when the prompt identification of pathogen may be crucial for a patient’s prognosis. Proteus spp. can be isolated from different material, including samples of food, feces, soil or polluted water, although most frequently detection of Proteus is performed in urine. Samples may be prepared by concentration of

10 ml urine by centrifugation at 2500 × g for 20 min at 4°C. The preparation of other samples (e.g., food, soil) usually requires a precedent step of homogenization in sterile PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4). If the amount of bacteria is low or there is a problem with the large number of contaminating bacteria, Proteus can be isolated from the samples after cultivation on selective and differential media (described in Section 30.1.4.1) within 24 h at 37οC. The swarming ability of Proteus rods can be inhibited by increasing concentration of agar in the medium or by use of one of a number of substances specified above. A well-isolated colony should be used for inoculation of liquid medium such as Luria-Bertani broth. For DNA isolation, place 1 ml of pure overnight culture in a sterile Eppendorf tube and centrifuge at 14000 × g for 1  min. Resuspend the pellet in 250 μl of Tris-EDTA (TE) buffer, add 50 μl of lysozyme in TE (10 mg/ml) and incubate for 15 min at 37°C. Add 60 μl of STEP buffer and incubate for 2 h at 50°C. Extract two to three times with equal volume of phenol:chloroform solution, each time collecting the aqueous phase into a new tube. Add equal volume of chloroform:isoamyl alcohol solution (24:1) and repeat extraction. Transfer the aqueous phase into a new tube, add 0.1 volume of 3 M sodium acetate, mix, and precipitate with 2.5 volumes of cold 96% ethanol, then centrifuge at 14000 × g for 10 min at 4°C. Dry the pellet and resuspend in 100 μl of TE buffer. Add 5 μl of RNaseA (5 mg/ml in TE), incubate for 30 min at 37°C. Extract once with chloroform:isoamyl alcohol solution and precipitate with ethanol. Wash the pellet with 1 ml of 70% ethanol, centrifuge, and dry. Resuspend in 100 μl of TE buffer and determine the concentration spectrophotometrically. Store at –20°C.

30.2.3  Detection Procedure Principle: As mentioned earlier, no particular method has been officially recommended for molecular detection of Proteus bacilli. Among different protocols presented in Table 30.3 the ones focused on the amplification of 16S rDNA region seem to be the most universal and are relatively simple to perform. There are several primer pairs in use which are anchored in the 16S rDNA conservative region which allow for the amplification of the variable regions of this gene. Here, we present a pair of universal primers targeting 3′ end of the gene which were designed originally by Lu et al.80 (U1: 5′-CCAGCAGCCGCGGTAATACG-3′; U2: 5′-ATCGG(C/T)TACCTTGTTACGACTTC-3′) and in case of Proteus give rise the 997 bp fragment. Procedure: (1) Prepare PCR mixture (50 μl) containing 5 μl of 10 × polymerase buffer, 200 μM of each deoxynucleoside triphosphate (dNTP), 0.2 μM of each primer, 20–50 ng of genomic DNA (prepare also a negative control in which deionized water should be used instead of DNA), and 2.5 U of Taq DNA polymerase. (2) Apply the following cycling program: one cycle of 94°C for 10 min; 35 cycles of 94°C for 1 min, 55°C

30.3 Conclusions and Future Perspectives Genus Proteus is a group of bacteria inhabiting environment, especially polluted water and soil, from which they can colonize the gastrointestinal tract of humans and animals. Among eight species belonging to Proteus, only P. myxofaciens is not involved in pathogenesis of mammals. The remaining seven species: P. mirabilis, P. vulgaris, P. penneri, P. hauserii, as well as genomospecies 4, 5, and 6 are able to develop a pathogenic process in their hosts, leading to several different diseases among which UTI are the most common. In clinical practice, diagnosis of Proteus is usually based on the classical microbiological methods, aimed to the detection of a very typical motile growth on agar media and followed by the analysis of specific biochemical properties, thus making the whole procedure relatively fast and reliable. However, under favorable conditions, especially within the group of patients with chronically infected urinary tracts, dissemination of these bacteria leading to septicemia may occur. In such cases, prompt identification of the infecting microorganism is crucial for the application of the targeted therapy, allowing

25 bp DNA ladder

P. mirabilis/HaeIII

P. vulgaris/HaeIII

P. mirabilis/MnlI

P. vulgaris/MnlI

P. mirabilis/HpaII

for 1 min and 72°C for 2 min; one cycle of 72°C for 10 min. (3) Check the size and the specificity of the PCR product by running 5 μl on the 1% agarose gel in TAE buffer, along with the negative control and the linear DNA ladder. (4) To sequence the PCR product, remove primers from the sample by using one of several available commercial kits for PCR product purification and send it to the sequencing facility. Compare the results using AlignX software with the sequences from the GenBank database (accession numbers: P. mirabilis, AJ301682; P. vulgaris, AJ233425; P. penneri, AJ634474). (5) Alternatively, analyze the PCR product using RFLP. Figure 30.2 presents restriction patterns obtained in silico using Vector NTI software after digestion with HaeIII, and MnlI endonucleases,80 as well as HpaII. Digestion with HaeIII gives genus specific pattern (34, 126, 161, 180, 217, and 279 bp), whereas MnlI (P. mirabilis: 14, 47, 65, 104, 119, 126, 157, 176, 189 bp; P. vulgaris: 14, 14, 47, 65, 104, 119, 126, 157, 175, 176 bp) and HpaII (P. mirabilis: 11, 81, 130, 130, 202, 443 bp; P. vulgaris: 11, 81, 130, 130, 137, 202, 306 bp) yield species specific bands. For digestion use 5 μl of PCR product, 2 μl of digestion buffer supplied by the enzyme manufacturer, 5–10 U of appropriate restriction endonuclease and fill up to 20 μl with deionized water. Incubate for 1 h at 37°C. Precipitate the sample with 96% ethanol, dry the pellet and suspend it in 10 μl of water. Run through the 7% polyacrylamide gel in TBE buffer along with the linear DNA ladder of the required range, stain with ethidium bromide and visualize under UV light.

1

2

3

4

5

6

P. vulgaris/HpaII

427

Proteus

7

500 450 400 350 300 250 200 175 150 125 100 75

50

25

Figure 30.2  RFLP analysis of 16S rDNA amplified fragment in 7% polyacrylamide gel.

for the fast eradication of the life-threatening agent. Thus, fast, reliable, and sensitive methods of molecular detection, identification, and typing of Proteus should be implemented into diagnostic protocols of microbiological laboratories. The methods based on the amplification of the specific regions followed by the analysis of the nucleotide sequence, although sensitive and very reliable when focused on the proper targets, cannot be commonly used when the possibility of multibacterial infection may occur. A pure population of bacteria is required for DNA isolation, hence their application needs the preliminary stage during which a time-consuming classical microbiological differentiation has to be done in

428

order to obtain monomicrobial cultures. Moreover, despite the increasing availability of the sequencing services, the procedure, should it be quick, requires in-house performance which is expensive and needs the assistance of well-trained personnel, thus limiting its application to the reference laboratories. Instead of sequencing, such techniques as RFLP, SSCP, DGGE, or real-time PCR which are cheaper, easier to perform, and which solve, at least partially, the problem of multispecies infections, may by utilized for identification of the amplified specific fragments. On the other hand, to be included into routine practice, these methods require laborious, multistage standardization, that would lead to the development of guidelines allowing results comparable between different microbiological laboratories. As in the case of other pathogens, the DNA microarray technology seems to be the future in the field of fast diagnostics and typing of Proteus originating from biological specimens as well as from food. The increasing popularization of the automated microchip readers accompanied by their decreasing cost, as well as the development of commercially available standardized microchips containing probes against vast ranges of particular genera, species or even strains, will create a chance to utilize this method even in smaller laboratories. Currently, the major drawback of DNA microchip technology is incomplete knowledge about the genome sequence of particular bacterial species, which complicates the development of specific probes for detection as well as primers for amplification of suitable regions. One has to remember, that sensitive detection and precise identification of microorganisms requires often the utilization of more than one probe and sometimes also amplification of more than one target region in order to avoid false results, should mutation occur in one of the analyzed regions. It is worth mentioning, that the first complete sequence of P. mirabilis genome was only recently published,110 P. penneri genome sequence is currently under investigation as part of the Human Microbiome Project (http://nihroadmap.nih.gov/hmp/) while P. vulgaris and the remaining species of this genus, have not yet been considered for complete sequencing.

References





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6. O’ Hara, M.C. Classification of Proteus vulgaris biogroup 3 with recognition of Proteus hauseri sp. nov. nom. rev. and unnamed Proteus genomospecies 4, 5, and 6. Int. J. Syst. Evol. Microbiol., 50, 1869, 2000. 7. Janda, J.M. and Abbot S.L. The genus Proteus. In The Enterobacteriaceae, 2nd ed. ASM Press, Washington, DC, 2006. 8. Philips, J.E. In vitro studies on Proteus organisms of animal origin. J. Hyg. Camb., 53, 26, 1955. 9. Burke, J.P. et al. Proteus mirabilis infections in hospital nursery traced to human carrier. New Eng. J. Med., 284, 115, 1971. 10. Graber, C.D. and Lincoln, A.F. Infantile diarrhoea in the Denver area: significance of the Proteus-Providencia organisms. Pediatrics, 16, 585, 1955. 11. Müller, H.E. Occurrence and pathogenic role of MorganellaProteus-Providencia group bacteria in human feces. J. Clin. Microbiol., 23, 404, 1986. 12. Müller, H.E. The role of Proteeae in diarrhea. Zbl. Bakteriol., 272, 30, 1989. 13. Von Holy, A., Holzapfel, W.H. and Dykes, G.A. Bacterial populations associated with Vienna sausage packaging. Food Microbiol., 9, 45, 1992. 14. Ababouch, L. et al. Identification of histamine producing bacteria isolated form sardine (Sardina pilchardus) stored in ice and at ambient temperature (25°). Food Microbiol., 8, 127, 1991. 15. Gonzalez-Rodriquez, M.-N. et al. Numbers and types of microorganisms in vacuum-packed cold-smoked freshwater fish at retail level. Int. J. Food Microbiol., 77, 161, 2002. 16. Lopez-Sabater, E.I. et al. Bacteriological quality of tuna fish (Thunnus thynnus) destined for canning: effect of tuna handling on presence of histidine decarboxylase bacteria and histamine level. J. Food Protect., 57, 318, 1994. 17. Youssef, H., El-Timawy, A.K. and Ahmed, S. Role of aerobic intestinal pathogens of fresh water fish in transmission of human diseases. J. Food Prot., 55, 739, 1992. 18. Fernandez-Delgado, M. et al. Occurrence of Proteus mirabilis associated with two species of Venezuelan oysters. Rev. Inst. Med. Trop. S. Paulo., 49, 335, 2007. 19. Dankor, E.S., Aning, K.G. and Quaye, J. Bacterial contamination of informally marketed milk in Ghana. Ghana Med. J., 41, 58, 2007. 20. Deetae, P. et al. Production of volatile aroma compounds by bacterial strains isolated form different surface-ripened French cheeses. Appl. Microbiol. Biotechnol., 76, 1161, 2007. 21. Senior, B.W. Proteus, Morganella, and Providencia. In Topley and Wilson’s Microbiology and Microbial Infections, Colier L., Ed. p. 1035. Arnold, London, UK, 1998. 22. Rashid, T. and Ebringer, A. Rheumatoid arthritis is linked to Proteus – the evidence. Clin. Rheumatol., 26, 1036, 2007. 23. Belas, R. The swarming phenomenon of Proteus mirabilis. ASM News, 58, 15, 1992 24. Gygi, D., et al. A cell surface polysaccharide that facilitates rapid population migration by differentiated swarm cell of Proteus mirabilis. Mol. Microbiol., 17, 1167, 1995. 25. Rather, P.N. Swarmer cell differentiation in Proteus mirabilis. Environment. Microbiol., 7, 1065, 2005. 26. Belas, R. Proteus mirabilis swarmer differentiation and urinary tract infection In Urinary Tract Infections, Molecular Pathogenesis and Clinical Management, Mobley, H.L.T. and Warren, J.W. Eds. p. 271. ASM Press, Washington DC, 1996. 27. Coker, C. et al. Pathogenesis of Proteus mirabilis urinary tract infections. Microbes Infect., 2, 1497, 2000.

Proteus 28. Mobley, H.L.T. Virulence of Proteus mirabilis. In Urinary Tract Infections, Molecular Pathogenesis and Clinical Management, Mobley, H.L.T. and Warren, J.W., Eds. p. 245. ASM Press, Washington DC, 1996. 29. Róz∙ alski, A., Sidorczyk, Z. and Kotełko, K. Potential virulence factors of Proteus bacilli. Microbiol. Mol. Biol. Rev., 61, 65, 1997. 30. Róz∙ alski, A. Molecular basis of the pathogenicity of Proteus bacteria. Adv. Clin. Exp. Med., 11, 3, 2002. 31. Murphy, C. and Belas, R. Genomic rearrangements in the flagellin genes of Proteus mirabilis. Mol. Microbiol., 31, 679, 1999. 32. Loomes, L.M., Senior, B.W. and Kerr, M.A. A proteolytic enzyme secreted by Proteus mirabilis degrades immunoglobulins of the immunoglobulin A1 (IgA1), IgA2, and IgG isotypes. Infect. Immun., 58, 1970, 1990. 33. Belas, R., Manos, J. and Suvanasuthi, R. Proteus mirabilis ZapA metalloprotease degrades a broad spectrum of substrates, including antimicrobial peptides. Infect. Immun., 72, 5159, 2004. 34. Wassif, C., Cheek, D. and Belas, R. Molecular analysis of metalloprotease from Proteus mirabilis. J. Bacteriol., 177, 5790, 1995. 35. Walker, K.E. et al. ZapA, the IgA-degrading metelloprotease of Proteus mirabilis, is a virulence factor expressed specifically in swarmer cells. Mol. Microbiol., 32, 825, 1999. 36. Loomes, L.M., Senior, B.W. and Kerr, M.A. Proteinases of Proteus sp. purification, properties, and detection in urine in infected patients. Infect. Immun., 60, 2267, 1992. 37. Jones, B.D., et al. Construction of urease-negative mutant of Proteus mirabilis: analysis of virulence in mouse model of ascending urinary tract infection. Infect. Immun., 58, 1120, 1990. 38. Dumanski, A.J., et al. Unique ability of Proteus mirabilis capsule to enhance mineral growth in infectious urinary calculi. Infect. Immun., 62, 2998, 1994. 39. Torzewska, A., Sta˛ czek, P. and Róz∙ alski, A. The crystallization of urine mineral components may depend on the chemical nature of Proteus endotoxin polysaccharides. J. Med. Microbiol., 52, 471, 2003. 40. Drechsel H.A., et al. α-Keto acids are novel siderophores in the genera Proteus, Providencia and Morganella and are produced by amino acid deaminases. J. Bacteriol., 175, 2727, 1993. 41. Welch, R.A. Identification of two different hemolysin determinants in uropathogenic Proteus isolates. Infect. Immun., 55, 2183, 1987. 42. Koronakis V. et al. The secreted hemolysins of Proteus mirabilis, Proteus vulgaris, and Morganella morganii are genetically related to each other and to the alpha-hemolysin of Escherichia coli. J. Bacteriol., 169, 1509, 1987. 43. Róz∙ alski A. and Kotełko K. Hemolytic activity and invasiveness in strains of Proteus penneri. J. Clin. Microbiol., 25, 1094, 1987. 44. Kwil, I., et al. unpublished data, 2006. 45. Peerbooms, P.G., Verweij, A.M.J.J. and MacLaren, D.M. Vero cell invasiveness of Proteus mirabilis. Infect. Immun., 4, 1068, 1984. 46. Róz∙ alski, A., Długon´ ska, H. and Kotełko, K. Cell invasiveness of Proteus mirabilis and Proteus vulgaris strains. Arch. Immunol. Ther. Exp., 34, 505, 1986. 47. Chippendale, G.R. et al. Internalisation of Proteus mirabilis by human renal epithelial cells. Infect. Immun., 62, 3115, 1994. 48. Allison, C. et al. Ability of Proteus mirabilis to invade urothelial cells is coupled to motility and swarming differentiation. Infect. Immun., 60, 4740, 1992.

429 49. Raetz, C.R. and Whitefield, C. Lipopolysaccharide endotoxin. Ann. Rev. Biochem., 71, 635, 2002. 50. Róz∙alski, A. Lipopolysaccharide (LPS, endotoxin) of Proteus bacteria – chemical structure, serological specificity and the role in the pathogenicity. Folia Biol. Oecol., 4,5, 2008 (accepted). 51. Hawkey, P.M. Proteus, Providencia, and Morganella spp. In Principles and Practice in Clinical Bacteriology, 2nd ed. Gillespie, S.H. and Hawkey P.M., Eds. John Wiley & Sons, Ltd. England, 2006. 52. Weissfeld, A.S. et al. Enterobacteriaceae. In Clinical and Pathogenic Microbiology, 2nd ed. Howard, B.J. et al., Eds. Mosby, st Louis, 1994. 53. Hawkey, P.M., Mc Cormick, A. and Simpson, R.A. Selective and differential medium for primery isolation of members of the Proteeae. J. Clin. Microbiol., 23, 600, 1986. 54. Urbanova, E. Selective medium for primery isolation of members of the tribe Proteeae. Folia Microbiol., 44, 629, 1999. 55. Senior, B.W. Media and tests to simplify the recognition and identification of members of the Proteeae. J. Med. Microbiol., 46, 39, 1997. 56. Müller, H.E. Proteus penneri showing a green colour reaction with Kovacs’ indole reagent. Zentbl. Bacteriol. Hug. A, 261, 198, 1986. 57. Farmer III, J.J. et al. Biochemical identification of new species and biogroups of Enterobacteriaceae isolated from clinical specimens. J. Clin. Microbiol., 21, 46, 1985. 58. Hickman, F.W. and Farmer III, J.J. Differentiation of Proteus mirabilis by bacteriophage typing and the Dines reaction. J. Clinical. Microbiol., 3, 350, 1976. 59. Senior, B.W. Typing of Proteus strains by proticine production and sensitivity. J. Med. Microbiol., 10, 7, 1977. 60. Senior, B.W. and Larsson, P. A highly discriminatory multityping scheme for Proteus mirabilis and Proteus vulgaris. J. Med. Microbiol., 16, 193, 1983. 61. Kauffmann, F. The Bacteriology of Enterobacteriaceae. The Williams & Wilkins, Co. Baltimore, MD, 1966. 62. Larsson, P. Serology of Proteus mirabilis and Proteus vulgaris. Methods Microbiol., 14, 187, 1984. 63. Penner, J.L., Hennessy J.N. Separate O-grouping schemes for serotyping clinical isolates of Proteus vulgaris and Proteus mirabilis. J. Clin. Microbiol., 12, 77, 1980. 64. Knirel, Y.A. et al. Structural study of O-specific polysaccharide of Proteus. J. Carbohydr. Chem., 12, 379, 1993. 65. Knirel, Y.A, et al. Structure of O-antigenic polysaccharides of Proteus bacteria. Pol. J. Chem., 73, 859, 1999. 66. Drzewiecka, D., Zych, K. and Sidorczyk, Z. Characterization and serological classification of a collection of Proteus penneri clinical strains. Arch. Immunol. Ther. Exp., 52, 121, 2004. 67. Bartodziejska, B. et al. Structural and serological studies on a new acidic O-specific polysaccharide of Proteus vulgaris O32. Eur. J. Biochem., 256, 488, 1998 68. Torzewska, A. et al. Structure of the O-specific polysaccharide from Proteus vulgaris O37 and close serological relatedness of the lipopolysaccharides of P. vulgaris O37 and P. vulgaris O46. FEMS Immunol. Med. Microbiol., 31, 227, 2001. 69. Torzewska, A. et al. Structure and serology of the O-antigen of Proteus strains classified into serogroup O17 and former serogroup O35. Arch. Immunol. Therap. Exp., 52, 277, 2006. 70. Cedzynski, M. et al. Structural and immunochemical studies of two cross-reactive Proteus mirabilis O-antigens, O6, and O23, containing β-1→3-linked 2-acetamido-2-deoxy-d-glucopyranose residues. Microbiol. Immunol., 42, 7, 1998.

430 71. Toukach, F.V. et al. Structure of a new acidic O-antigen of Proteus vulgaris O22 containing O-acetylated 3-acetamido3,6-dideoxy-d-glucose. Carbohydr. Res., 318, 146, 1999. 72. Kaca, W. et al. Structure of the O-specific polysaccharide of Proteus mirabilis S 1959. Arch. Immunol. Ther. Exp., 35, 431, 1987. 73. Shashkov, A.S. et al. Structures of new acidic O-specific polysaccharides of the bacterium Proteus mirabilis serogroups O26 and O30. FEBS Lett., 386, 247, 1996. 74. Radziejewska-Lebrecht, J. et al. Structure and epitope characteristic of O-specific polysaccharide of Proteus mirabilis O28 containing amides of d-galacturonic acid with l-serine and l-lysine. Eur. J. Biochem., 230, 705, 1995. 75. Arbatsky, N.P., et al. Structure of the O-specific polysaccharide of Proteus mirabilis O11, another Proteus O-antigen containing an amide of d-galacturonic acid with l-threonine. Carbohydr. Res., 323, 81, 2000. 76. Kocharova, N.A. et al. Structure of the O-specific polysaccharide of Providencia rustigianii O14 containing Nε-[(S)-1carboxyethyl]-Nα-(D-galacturonoyl)-L-lysine. Carbohydr. Res., 338, 1009, 2003. 77. Torzewska, A. et al. Serological characterization of the O-specific polysaccharide of Providencia alcalifaciens O23. Arch. Immunol. Therap. Exp., 52, 43, 2004. 78. Kocharova, N.A. et al. Structure of the O-polysaccharide of Providencia stuartii O4 containing 4-(N-acetyl-L-aspart-4-yl) amino-4,6-dideoxy-D-glucose. Carbohydr. Res., 339, 195, 2004. 79. Torzewska, A. et al. Structure of the O-polysaccharide of Providencia stuartii O33 containing 4‑(N‑acetyl-d-aspart4-yl)amino-4,6-dideoxy-d-glucose. FEMS Immunol. Med. Microbiol., 41, 133, 2004. 80. Lu, J.J. et al. Use of PCR with universal primers and restriction endonuclease digestions for detection and identification of common bacterial pathogens in cerebrospinal fluid. J. Clin. Microbiol., 38, 2076, 2000. 81. Vliegen, I. et al. Rapid identification of bacteria by realtime amplification and sequencing of the 16S rRNA gene. J. Microbiol. Methods, 66, 156, 2006. 82. Widjojoatmodjo, M.N., Fluit, A.C., Verhoef, J. Molecular identification of bacteria by fluorescence-based PCR-singlestrand conformation polymorphism analysis of the 16S rRNA gene. J. Clin. Microbiol., 33, 2601, 1995. 83. Schabereiter-Gurtner, C. et al. 16S rDNA-based identification of bacteria from conjunctival swabs by PCR and DGGE fingerprinting. Invest. Ophthalmol. Vis. Sci. 42, 1164, 2001. 84. Mansy, M.S. et al. Amplification of Proteus mirabilis chromosomal DNA using the polymerase chain reaction. Mol. Cell. Probes., 13, 133–40, 1999. 85. Takeuchi, H. et al. Detection of Proteus mirabilis urease gene in urinary calculi by polymerase chain reaction. Int. J. Urol., 3, 202, 1996. 86. Kwil, I. et al. Use of randomly amplified polymorphic DNA (RAPD) analysis for identification of Proteus penneri. Adv. Exp. Med. Biol., 485, 321, 2000. 87. Kessler, C. and Manta, V. Specificity of restriction endonucleases and DNA modification methyltransferases–a review (Edition 3). Gene, 16, 1, 1990. 88. Roberts, R.J. Restriction enzymes and their isoschizomers. Nucleic Acids Res., 18 Suppl., 2331, 1990. 89. Lee, K.F. et al. Restriction endonucleases in clinical isolates of Shigella spp. J. Med. Microbiol., 46, 949, 1997. 90. Stephenson, F.H. et al. Comparison of the nucleotide and amino acid sequences of the RsrI and EcoRI restriction endonucleases. Gene, 85, 1, 1989.

Molecular Detection of Foodborne Pathogens 91. Metherell, L.A., Hurst, C. and Bruce, I.J. Rapid, sensitive, microbial detection by gene amplification using restriction endonuclease target sequences. Mol. Cell. Probes, 11, 297, 1997. 92. Versalovic, J., Koeuth, T. and Lupski, J.R. Distribution of repetitive DNA sequences in eubacteria and application to fingerprinting of bacterial genomes. Nucleic Acids Res., 19, 6823, 1991. 93. Serwecin´ ska, L. et al. Genomic fingerprinting of Proteus species using repetitive sequence based PCR (rep-PCR). Acta Microbiol. Pol., 47, 313, 1998. 94. Hong, B.X. et al. Application of oligonucleotide array technology for the rapid detection of pathogenic bacteria of foodborne infections. J. Microbiol. Methods, 58, 403, 2004. 95. Jin, L.Q. et al. Detection and identification of intestinal pathogenic bacteria by hybridization to oligonucleotide microarrays. World J. Gastroenterol., 11, 7615, 2005. 96. Liao, J.C. et al., Use of electrochemical DNA biosensors for rapid molecular identification of uropathogens in clinical urine specimen. J. Clin. Microbiol., 44, 561, 2006. 97. Kostic´, T. et al. A microbial diagnostic microarray technique for the sensitive detection and identification of pathogenic bacteria in a background of nonpathogens. Anal. Biochem., 360, 244, 2007. 98. Grimont, F. et al. rRNA gene restriction patterns of Legionella species: a molecular identification system. Res. Microbiol., 140, 615, 1989. 99. Irino, K. et al. rRNA gene restriction patterns of Haemophilus influenzae biogroup aegyptius strains associated with Brazilian purpuric fever. J. Clin. Microbiol., 26, 1535, 1988. 100. Tamayo, M. et al. Molecular epidemiology of Vibrio cholerae O1 isolates from Colombia. J. Med. Microbiol., 46, 611, 1997. 101. Kennett, C.A. and Stark, B. Automated ribotyping for the identification and characterization of foodborne clostridia. J. Food Prot., 69, 2970, 2006. 102. Voskressenskaya, E. et al. Evaluation of ribotyping as a tool for molecular typing of Yersinia pseudotuberculosis strains of worldwide origin. J. Clin. Microbiol., 43, 6155, 2005. 103. Pignato, S. et al. Molecular characterization of the genera Proteus, Morganella, and Providencia by ribotyping. J. Clin. Microbiol., 37, 2840, 1999. 104. Pfaller, M.A. et al. Evaluation of the discriminatory powers of the Dienes test and ribotyping as typing methods for Proteus mirabilis. J. Clin. Microbiol., 38, 1077, 2000. 105. Sabbuba, N.A., Mahenthiralingam E., Stickler D.J. Molecular epidemiology of Proteus mirabilis infections of the catheterized urinary tract. J. Clin. Microbiol., 41, 4961, 2003. 106. Bingen, E. et al. Arbitrarily primed polymerase chain reaction provides rapid differentiation of Proteus mirabilis isolates from a pediatric hospital. J. Clin. Microbiol., 31, 1055, 1993. 107. Cies´likowski, T. et al. Tandem tetramer-based microsatellite fingerprinting for typing of Proteus mirabilis strains. J. Clin. Microbiol., 41, 1673, 2003. 108. Kowalczyk, M. and Sidorczyk, Z. Determination of genetic diversity of Proteus penneri strains using rep-PCR. Adv. Exp. Med. Biol., 485, 315, 2000. 109. Drzewiecka, D. et al. Structure and serological properties of the O-antigen of two clinical Proteus mirabilis strains classified to a newly created Proteus O77 serogroup. FEMS Immunol. Med. Microbiol., 54, 185, 2008. 110. Pearson, M.M. et al. The complete genome sequence of uropathogenic Proteus mirabilis, a master of both adherence and motility. J. Bacteriol., 190, 4027, March 28, 2008.

31 Pseudomonas

Olga Zaborina and John Alverdy University of Chicago

Contents 31.1 Introduction.....................................................................................................................................................................431 31.1.1 Pseudomonas aeruginosa is a Foodborne Pathogen.........................................................................................431 31.1.2 Risk of Acquisition of P. aeruginosa in Hospitalized Patients........................................................................ 432 31.1.3 Intestinal Pathogenesis of P. aeruginosa.......................................................................................................... 434 31.1.4 Novel Nonantibiotic Strategies to Contain Intestinal P. aeruginosa................................................................ 436 31.1.5 Molecular Tools for P. aeruginosa Detection................................................................................................... 436 31.1.5.1 P. aeruginosa Detection in Food Products...................................................................................... 436 31.1.5.2 Detection of P. aeruginosa in Hospitals Clinical Laboratories....................................................... 439 31.1.5.3 Assessment of Virulence Potential in P. aeruginosa....................................................................... 439 31.2 Methods.......................................................................................................................................................................... 440 31.2.1 Sample Preparation........................................................................................................................................... 440 31.2.2 Detection Procedure......................................................................................................................................... 440 31.3 Conclusion...................................................................................................................................................................... 440 References.................................................................................................................................................................................. 440

31.1  Introduction 31.1.1 Pseudomonas aeruginosa is a Foodborne Pathogen The genus Pseudomonas comprises a number of Gramnegative, aerobic, rod-shaped free-living bacterial species belonging to the phylum Proteobacteria, class Gamma Proteobacteria, order Pseudomonadales, family Pseudomonadaceae. Being a most notable Pseudomonas species, Pseudomonas aeruginosa is traditionally considered to be a human opportunistic pathogen that infects the lungs of patient with the genetic disorder cystic fibrosis and the wounds of burn patients. P. aeruginosa is a foodborne pathogen. Although the role of P. aeruginosa as a foodborne pathogen is often not strictly defined, there is a growing body of evidence to suggest that its primary site of colonization in lungs and other infections is the gastrointestinal tract where it presents in up to 10% of healthy patients. Even in patients with cystic fibrosis, acquisition of P. aeruginosa via the oral route is postulated to precede lung infection either as a result of direct gastric to airway contamination or via hematogenous spread.1–6 Thus, although P. aeruginosa it is not listed in “Foodborne Pathogenic Microorganisms and Natural Toxins” Handbook (http://www.cfsan.fda.gov/~mow/intro.html) produced by FDA, or among Foodborne Pathogens of Animal Origin,7 its presence in contaminated food and its primary site of colonization being the intestine beg a reconsideration of this pathogen to be classified as a foodborne pathogen. Although P. aeruginosa can be present or become colonized within the

intestinal tract reservoir, it is considered to be opportunistic in that most case reports where it is described as causing either diarrhea, intestinal necrosis, or widespread gut-derived septicemia, involve patients with significant immunocompromises including burn injury, organ transplantations, AIDS, radiation, or steroid use. P. aeruginosa is the most common organism causing lethal gut-derived sepsis following bone marrow transplantation and radiation therapy. Yet there are multiple reports documenting that even in healthy subjects, P. aeruginosa can cause a syndrome of diarrhea, fever, and lethal sepsis from intestinal dissemination such as in children of age up to 2 years old.8–12 Cases including ecthyma gangrenosum,13 characteristic skin lesion,14,15 neutropenia,12 and lethal sepsis have been described in previously healthy children,8,9 and have been shown to be prevalent in boys, in infants, and during warm weather seasons.8 In summary, P. aeruginosa is prevalent in the human intestinal during both health and illness and can cause lethal sepsis (herein termed “lethal gut-derived sepsis”) from within this site by mechanisms that remain to be elucidated. Whether lethal gut-derived sepsis due to P. aeruginosa is strain specific, specific to local environmental condition, or a host specific phenomenon, is not fully understood. Its importance as a foodborne pathogen is underscored by its ubiquitous presence in the environment. Environmental sources of P. aeruginosa. In native environments, P. aeruginosa is commonly found in soil,16 spring water,17 river water,18 artesian water,19 ocean water,20 on the surfaces of plants21,22 and animals.23,24 In many cases, environmental reservoirs of P. aeruginosa originate from human activities such as river water downstream from urban areas 431

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contaminated with pig feces,25 mushroom-growing fields,21,26 technical water systems,27 gasoline-contaminated aquifer,28 and contaminated soils.29,30 The urban environment and its attendant concentration of human activity increase the risk of human exposure to P. aeruginosa. For example, P. aeruginosa at high cell densities of 104 –107 CFU/g have been found in mushrooms purchased from several supermarkets, originally grown in five different countries, as well as in packaged bean sprouts.21 Selective advantage for P. aeruginosa in human activity environments. The use of man made chemicals including poorly degradable pesticides may contribute to selective pressures on the natural microflora that provide a temporary selective advantage for P. aeruginosa that becomes resistant to many of these chemicals allowing it to persist when other microflora are eliminated31–34 and can utilize these compounds as a source of carbon and energy.34–39 Acquisition of antibiotic resistance in P. aeruginosa in human activity environments. To be protected from chemical stress, efflux pumps are activated in P. aeruginosa to excrete these toxic compounds. For example, a common environmental contaminant is polychlorinated aromatic compound pentachlorophenol, an extremely recalcitrant compound commonly used as a pesticide and wood preservative.40,41 In response to this anthropogenic chemical, the multidrug efflux pump MexAB-OprM was found to be upregulated in P. aeruginosa to protect it from the toxic effect of pentachlorophenol.42 Notably, overexpression of MexAB-OprM efflux pump accompanies multiple antibiotic resistance in P. aeruginosa43–45 that can lead to the selection of multi-drug resistant (MDR) environmental strains. Indeed, environmental isolates (mainly from oil-contaminated soils) have been found to contain multidrug resistance determinants and can actively efflux synthetic antibiotics.46 Finally, P. aeruginosa is able to consume antibiotics as a sole source of carbon and energy. Recently, bacteria classified as of 11 orders have been isolated from various soil samples that were able to grow in media containing only antibiotics as carbon source. Among them, Burkholderiades and Pseudomonadales represent the major orders.47

31.1.2 Risk of Acquisition of P. aeruginosa in Hospitalized Patients Gastrointestinal colonization with P. aeruginosa is common in hospitalized patients. Hospital environments are a major source of intestinal contamination with P. aeruginosa as the pathogen is present in a variety of sources of drinking water, food, feeding tubes, and surgical instruments that directly contact the intestinal tract.5,19,48–52 Several prospective studies have documented that patients culture positive for P. aeruginosa in rectal and oropharyngeal samples both upon admission and during hospitalization.53–57 Up to 20% of fecal samples were positive for P. aeruginosa upon admission to the hospital, and up to 50% became culture positive within a week of hospitalization.53,54,58 The highest prevalence of intestinal carriage of P. aeruginosa has been noted for patients

Molecular Detection of Foodborne Pathogens

in the Surgery Intensive Care Units (ICUs).53,54,58–61 As such, both endogenous and exogenous sources remain important mechanisms by which P. aeruginosa cross-transmission persists as a major problem in hospitals. Finally, the finding that hospital foods including fresh vegetables and salads are a source of intestinal colonization by P. aeruginosa, provide evidence hospitalized patients are exposed to this pathogen via multiple sources.50,62–66 Impact of antibiotic use on P. aeruginosa gastrointestinal colonization in hospitalized patients. The promiscuous use of antibiotics in hospitalized patients remains an unavoidable but not inconsequential practice that has led to the emergence of the concept of suprainfection. Loss of the protective intestinal flora as a result of this practice, coupled with exposure to hospital pathogens from various sources, results in rapid colonization by highly opportunistic pathogens such as P. aeruginosa,67–69 Candida albicans,70,71 Staphylococcus auerus,72 and Streptococcus faecalis.73,74 Several studies have documented a direct relationship between antibiotic use and emergence of infections caused by P. aeruginosa.75–77 A significant association between fluoroquinolone, imipenem, and ceftazidime use, and intestinal (GI) colonization by P. aeruginosa strains that develop resistance to these drugs has been documented.59,67 As P. aeruginosa is intrinsically resistant to a large number of antimicrobial drugs, only a limited list antibiotics remain that are effective for its treatment that include: carbapenems (imipenem), cephems (ceftazidime, cefoperazone), aminoglycosides (amikacin, tobramycin, gentamicin), fluoroquinolones (ciprofloxacin), and β-lactame/βlactamase inhibitor combinations (piperacillin/tazobactam).78 Antimicrobial resistance to P. aeruginosa has emerged as an important determinant of outcome during infection.67,79–81 World-wide infections due to antibiotic resistant organisms are most prevalent in hospital ICUs,82 where P. aeruginosa has occupied the top position, a position that has remained unchanged for decades.62–64,83,84 Due to the promiscuous use of antibiotics in this group of patients, within as little as 3 days in an ICU, the feces of more than 50% of patients in an ICU become culture positive for P. aeruginosa with up to 30% of these strains being antibiotic resistant.57 These findings underscore the remarkable exposure of the sickest hospitalized patients to P. aeruginosa despite highly vigilant protocols to limit its spread from patient to patient. Antibiotic restriction policy. Since the overuse of antibiotics in ICUs can lead to an increase in the recovery of antibiotic resistant strains causing infection, several hospitals have instituted an antibiotic restriction policy to limit the development and dissemination of resistant microorganisms. While this policy has been successful in suppressing specific outbreaks of infection with certain antibiotic-­resistant bacteria, overall it has been difficult to demonstrate the effect of this policy on preventing the development of antibiotic resistance in other competing strains. For example, in 1993, restriction of fluoroquinolone use in the ICU of Bellevue Hospital (France) over a 5-month period resulted in the reestablishment of sensitivity of P. aeruginosa to fluoroquinolones. However, after terminating the program, a high

Pseudomonas

percentage (71.3%) of P. aeruginosa developed resistance to ciprofloxacin in the same ICU.85 Similarly, a restriction policy in the ICU in Greece to the empirical use of fluoro­ quinolones and ceftazidime86 for 18 months led to a significant decrease in use of ceftazidime (92.5% decrease) and quinolones (55.4% decrease) associated with a significant increase in susceptibility of P. aeruginosa to ciprofloxacin and ceftazidime. Yet concomitant with this shift was an increase in the use of cefepime (270%) and colistin, and a significant decrease in the susceptibility of P. aeruginosa to carbapenem and meropenem.86 In another study, restricted use of cephalosporins led to a 44.0% reduction in infections due to Klebsiella within a 1-year observation period that inadvertently resulted in a 140.6% increase in the use of imipenem. However the shift from cepahlosporins to imipenem during the 1-year observation period led to an 68.7% increase in imipenem-resistant P. aeruginosa throughout the medical center87 questioning the overall effectiveness of this strategy. In another approach, antibiotic class cycling (or antibiotic rotation), was also shown to be of limited value as a sole intervention.88 Under this approach, two or more antibiotic classes are alternated on a time scale of months to years reasoning that when resistance to one class reaches high frequency, a scheduled switch to another class of antibiotic will promote susceptibility to a new drug,88–90 thus delaying the development and spread of antibiotic resistance in hospitals. Although this approach sounds promising as a long-term strategy, it has been shown to be ineffective in several intervention trials that either failed to show any advantage to cycling24 or only showed a trend to a possible advantage.91–93 A mathematical model of antimicrobial cycling in a hospital setting has been recently developed to explore the efficacy of cycling programs.94 This model demonstrated that heterogeneous antibiotic use slows the spread of resistance, however at the scale relevant to bacterial populations, heterogeneity of bacterial populations is greater than antibiotic heterogeneity performed by cycling. As a consequence, cycling is unlikely to be effective and may even hinder resistance control measures.94 Strategies incorporating multiple interventions, including antibiotic heterogeneity, shorter courses of treatment and narrowing or de-escalation of antibiotics on the basis of culture results, appear to be more effective88 but still do not solve the problem of antibiotic resistance. Emergence of MDR in P. aeruginosa. In the last two decades, an increase in the frequency of MDR strains of P. aeruginosa has been reported. MDR P. aeruginosa is defined as resistance to at least three drugs in the following classes: β-lactams, carbapenems, aminoglycosides, and fluoroquinolones.95 Risk factors for MDR P. aeruginosa infection included prolonged hospitalization, exposure to antimicrobial therapy, and an immunocompromised state. Emergence of multi-drug antibiotic resistance during antibiotic therapy has been reported in 27–72% of patients with initially susceptible P. aeruginosa isolates.96 The most effective strategy of MDR P. aeruginosa treatment includes the combination of colistin (old polymixin with a narrow antibacterial spectrum

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covering aerobic Gram-negative bacilli) with β-lactams, ciprofloxacin or rifampicin96–103 although colistin carries a high risk of toxicity. Does multi-antibiotic resistance confer virulence in P. aeruginosa? Although the increasing prevalence of MDR strains of P. aeruginosa among critically ill humans is of significant concern, the evidence that multi-antibiotic resistance itself confers a more virulent phenotype is highly variable.104 A recent analysis of consecutively isolated MDR strains of P. aeruginosa from critically ill hospitalized patients that assessed the virulence of a given strain against the intestinal epithelium using cultured Caco-2 cells, demonstrated an extremely polarized array of phenotypes from nonmotile and nonadhesive to highly motile, adhesive, and disrupting intestinal epithelial monolayers.104 Decreased virulence of gentamicin resistant P. aeruginosa was demonstrated in rats and mice injected intraperitoneally with GmR and GmS strains.105 Similarly, at the Mayo Clinic, between the years 1970 and 1976, the virulence of gentamicin-resistant isolates appeared less than that of susceptible organisms, with bacteremia due to these isolates occurring in only three cases.106 Another report that analyzed 47 clinical antibiotic resistant strains of P. aeruginosa confirmed that the antibiotic resistance of P. aeruginosa is not generally associated with enhanced production of virulence factors.107 A possible explanation to these observations, with specific regards to P. aeruginosa might be that multi-drug antibiotic resistance strains are attenuated in virulence as a result of the overproduction of MDR efflux pumps MexAB-OprM, MexEF-OprN, and MexXY.108 The implication of efflux pumps activation to the resistance of P. aeruginosa to different classes of antibiotics such as fluoroquinolones, β-lactams, and aminoglycosides was widely demonstrated.109–113 MDR pumps can extrude antibiotics and therefore prevent their bactericidal effect; however, the same MDR pumps promote the efflux of quorum sensing signaling molecules114,115 thus preventing activation of quorum sensing. Unexpectedly, the overproduction of specific efflux pumps, either MexCD-OprJ or MexEF-OprN, has been shown to be associated with a reduction in the transcription of the type III secretory system due to the lack of expression of the exsA gene, encoding a central regulator of this machinery.108 The type III secretion system whose activation is induced upon contact with mammalian cells or at low calcium concentration in media, is responsible for the injection of virulent effector proteins directly in the cytoplasm of target cells through a needle-like structure.116–119 Since MDR pumps can extrude a wide range of compounds belonging to different structural families, it is possible that a yet-unknown intracellular signal exists that might allow ExsA to activate transcription of its regulated genes, that can be extruded by Mex pumps. Overexpression of efflux pumps in conjunction with the acquisition of MDR and down regulation of type III secretion system has been shown to lead to reduced bacterial virulence in P. aeruginosa in different model systems including Caenorhabditis elegans,120 Dictyostelium discoideum,121 and the macrophage cell line J774.108 Yet conversely, several reports have demonstrated that MDR P. aeruginosa are fully

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pathogenic104,109,122 by virtue of their ability to express MDR via various mechanisms without losing their ability to generate severe bloodstream infections109 and by their ability to display a highly adherent and disruptive phenotype against epithelial cells.104 Such a barrier disrupting phenotype in MDR P. aeruginosa clinical strains isolated from critically ill patients has been demonstrated using cultured intestinal epithelial cells and a mouse model of lethal intestinal sepsis due to P. aeruginosa following surgical injury.122 The virulence of MDR stains of P. aeruginosa in these models involved the expression of novel appendages in P. aeruginosa that were rich in the phosphate scavenging protein PstS. PstS rich appendages were induced during phosphate limitation and completely suppressed in phosphate-rich media. Injection of MDR strains directly into the intestinal tract of hepatectomy stressed mice, a surgical injury known to be accompanied by the phosphate depletion in intestinal mucus, caused high mortality rates (60–100%),123,124 while oral supplementation of phosphate in this model completely prevented mortality. Given the high incidence of acute depletion of phosphate during critical illness and major surgery,125–127 it is possible that prolonged exposure of MDR P. aeruginosa to a low phosphate microenvironment promotes mutations that lead to the expression of the phosphate signaling/scavenging system. As such, mutations might increase P. aeruginosa virulence expression as they acquire resistance genes to adapt and survive in harsh colonization niches such those encountered in the intestinal tract of a critically ill patient.

31.1.3 Intestinal Pathogenesis of P. aeruginosa The intestinal tract of a critically ill human exposes P. aeruginosa to a unique microenvironment in which to adapt and survive. The ecologically diverse and competitive environment of the mammalian intestinal tract is designed to impose a degree of colonization resistance to transient pathogens that maintains its stability and function. Under conditions of health where a diverse flora and intact immune system exist, home filed advantage and a co-evolved immune response prevent the colonization of pathogenic organism that are not part of the commensal flora. In this regard, bacteria themselves are part of the innate immune system of the intestinal tract as they maintain homeostasis on multiple levels including educating the immune system. This is in contradistinction to sterile organs such as the lung or kidney where colonization by bacteria is not present. Due to the high microbial burden in the mammalian gut, a thick protective mucus layer, especially in the colon, has formed to allow for the colonization of bacteria that remain physically distanced from the epithelial surface. Thus an innate mechanism of immune exclusion exists that allows for an anti-inflammatory mechanism to eliminate or tolerate nondangerous food, antigens, and commensal microorganisms (oral, mucosal tolerance).128 The apical side of the intestinal epithelium is highly resistant to various toxic and cytolytic bacterial exoproducts including those of P. aeruginosa exotoxin A and elastase,123,124,129 which is not the case in lung epithelium.130–133 As such, findings in

Molecular Detection of Foodborne Pathogens

lung and other organs in response to P. aeruginosa infection cannot be directly extrapolated to how inflammation proceeds in the intestine when confronted by exposure to this pathogen. During physiologic stress, tissue injury and critical illness, the intestinal microflora and the luminal microenvironment are drastically altered by multiple factors including antibiotic use, the release of local host tissue antibiotics and antimicrobial factors, a shift in pH and oxygen, and specific fuels upon which many microbes depend such as volatile fatty acids, phosphate, and iron. Mucus is also precipitously and significantly depleted during these disorders and their treatments. As the gut becomes further disabled by these physiologic disturbances, care givers must maintain host nutrition via highly unphysiologic routes including intravenous nutrition. Precisely how microbes adapt and survive this historically unprecedented environment from the standpoint of these selective pressures over the evolutionary time of a microbe has not been studied. Host stress shifts P. aeruginosa to a lethal phenotype. Organisms that colonize the intestinal tract of hospitalized patients represent a class or organisms referred to as nosocomial pathogens. The mechanisms by which these wily and resistant classes of organisms are able to colonize the intestinal tract are multifactorial and involve loss of the normal probiotic flora from antibiotic use, disruption of the innate immune system due to shock, immunosuppression, drugs, and rampant exposure to host stress derived factors in nutrient depleted local intestinal microenvironment. Opportunistic pathogens such as P. aeruginosa also may carry/upregulate resistance and virulence genes as they have had to adapt and survive multiple complex host environments as they pass from one host to another. The presence of P. aeruginosa in the human intestinal tract when the host is physiologically stressed or immunocompromised can cause severe sepsis syndrome from either translocation and dissemination of the organism across the intestinal wall into the systemic circulation, or can cause a syndrome of gut-derived sepsis whereby the bacteria is able to cause a lethal toxicosis from within the intestinal tract by disrupting the intestinal epithelial barrier to its lethal cytotoxins such as exotoxin A or elastase.134–137 Recent work in this area has advanced the concept that opportunistic pathogens colonizing a physiologic host can shift their phenotype from indolent colonizer to lethal pathogen in direct response to host tissue derived compounds that are released during tissue injury and stress.123 In the case of P. aeruginosa this shift can result in its transformation to an adhesive phenotype that disrupts intestinal epithelial integrity causing activation of a systemic inflammatory response, multiple organ dysfunction, and even death.2,123,124,129,138–143 For example, it has been demonstrated that following surgical injury, host tissue compounds are released into the intestine that activate the sense and response circuitry of P. aeruginosa such that it is transformed to express a virulence phenotype in response to specific environmental cues.129,139,142–144 When mice are subjected to a 30% surgical hepatectomy or ischemia/reperfusion of the small intestine and exposed to intestinal P. aeruginosa, lethality occurs as a result of a shift

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in the phenotype of P. aeruginosa to that of an adhesive and lethal strain.123 How intestinal P. aeruginosa causes systemic inflammation and organ damage without translocation and dissemination to remote organs. In working with the model of surgical injury, including a 30% surgical hepatectomy, or imposing a period of ischemia/reperfusion injury to the intestine, it has been discovered that P. aeruginosa expresses quorum sensing dependent virulence activation in direct response to soluble compounds released into the gut during these injuries.138,139 It has been further found that filtered intestinal contents isolated from surgically injured mice activate independent cultures of P. aeruginosa to express quorum sensing dependent virulence activation.139,142 It has been discovered that a particular adhesin, the PA-I lectin has been expressed following exposure to intestinal contents following surgical hepatectomy and that this glycoprotein plays a key role in the lethality of this organism when present in the intestinal tract of a stressed host. It has been discovered that the PA-I lectin dysregulates several intestinal epithelial tight junction proteins such as occludin and ZO-1 creating a epithelial permeability defect to lethal toxins of P. aeruginosa including elastase and exotoxin A. In fact, simultaneous injection of PA-I and exotoxin A into the mouse cecum during surgical hepatectomy resulted in death due to sepsis where injection of either compound alone had no effect.124 Systemic injection of PA-I had no effect, whereas injection of exotoxin A was lethal. Therefore after several studies in intestinal epithelial cells and mouse models, compelling evidence suggests that following surgical injury, P. aeruginosa is cued to express the quorum sensing dependent virulence determinant PA-I lectin which alters intestinal permeability resulting in penetration of bacterial cytotoxins into systemic compartment and death in mice. This process appeared to occur independent of bacterial dissemination, as both in control mice (mice subjected to sham laparotomy and intestinal inoculation with P. aeruginosa) and surgical injury (mice subjected to 30% surgical hepatectomy and intestinal inoculation with P. aeruginosa) bacteremia developed but did not correlate with mortality. As further evidence for this notion, when the same inoculum size of P. aeruginosa administered intravenously or intraperitoneally, no death has resulted, indicating that dissemination alone does not cause mortality or even systemic inflammation. Similar findings have been reported by others in both an intestinal inoculation model of P. aeruginosa as well as a model in rabbits in which the lung was inoculated. Therefore, P. aeruginosa virulence is activated locally and can cause systemic inflammation and death without translocation and dissemination by mechanisms that involve disruption of the barrier function of the epithelium to lethal cytotoxins. Yet the mechanisms by which local P. aeruginosa causes lethality are distinct depending on the tissue site. The lung is a sterile organ and normally is highly responsive to the presence of any bacteria. For example, exposure of lung cells to exotoxin A or elastase causes severe epithelial inflammation and loss of barrier130,132,133 function whereas this effect is not observed with cultured intestinal

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epithelial cells.123,124 Therefore, extrapolation of mechanisms from one site to another is not always possible. The specific environmental cues within the intestinal tract following surgical injury or physiologic stress, that are released into sites of P. aeruginosa colonization which activate its virulence have been identified. In various models of stress, interferon gamma, opioids (dynorphin, morphine), and end products of ischemia (adenosine) have been found to activate quorum sensing dependent virulence expression in P. aeruginosa. For example, interferon gamma binds to the outer membrane porin OprF that activates the quorum sensing circuitry of P. aeruginosa to increase PA-I and pyocyanin expression. In experiments with cultured intestinal epithelial cells, exposure of P. aeruginosa to interferon gamma shifts its phenotype to disrupt epithelial barrier function.143 Similar quorum-sensing effecting mechanisms have been demonstrated with the opioid dynorphin that is released from intestinal tissues in response to ischemic stress. Dynorphin binds to MvfR, a global transcriptional regulator of the quorum sensing circuitry of P. aeruginosa, and induces the expression of MvfR-regulated genes leading to enhanced production of pyocyanin and other compounds that alter the virulence of P. aeruginosa.142 Finally, adenosine, an epithelial cytoprotectant released by intestinal epithelial cells in the response to stress and ischemia, is taken up into the cytoplasm of P. aeruginosa, converted to inosine by P. aeruginosa adenosine deaminase, and inosine then activates the expression of the PA-I lectin causing intestinal epithelial barrier disruption.139 Taken together, these studies demonstrate how P. aeruginosa can intercept host stress signals, incorporate them into its virulence circuitry and mount a toxic offensive against the host without translocation and dissemination. Thus, in many circumstances, intestinal P. aeruginosa may remain clinically undetectable yet can cause sepsis in immunocompromised patients. Yet its presence in the intestinal tract alone is insufficient evidence that it is the cause of sepsis. As assays of its virulence phenotype are difficult to perform without reculturing the organism in nutrient rich and neutral environments, its relative virulence in a given patients has not been used as a measure of its potential threat. Because P. aeruginosa is so often recovered from the feces of hospitalized patients, its presence in this site cannot alone be used as an indicator of need for eradication or as a cause of sepsis. In addition, in hospitalized patients dying of sepsis, on in 50% of cases is the offending pathogen and original site of infection unknown. As autopsy studies of patient dying of sepsis indicate that the intestinal tract is a common site of microbial pathogenesis and ultimately death, a more complete understanding of the phenotypes and behavior of colonizing strains is needed. How intestinal P. aeruginosa causes systemic inflammation and organ damage via translocation and dissemination to remote organs. Animal models and human studies propose the traditional view on the mechanism by which intestinal P. aeruginosa causes host damage by offering evidence that it disseminates to remote organs causing bacteremic sepsis. In clinical studies, heavy growth of intestinal P.  aeruginosa is most often associated with diarrhea.

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This usually occurs in elderly and physiologically compromised hosts. In such cases there is rarely systemic symptomotology and there is little evidence that ongoing intestinal inflammation is present. In patients with AIDS, intestinal P. aeruginosa is a frequent cause of chronic diarrhea and the most common cause of death due to bacterial induced sepsis. Yet the most problematic causes of intestinal P. aeruginosa infections arise in those patients undergoing radiation injury, chemotherapy, or bone marrow transplantation. In such cases, the normal intestinal flora are disrupted or eradicated by the radiation effect itself, use of antibiotics, erosion of the nutritive mucus required for successful colonization, epithelial damage and desquamation, etc. In such cases, intestinal P. aeruginosa colonization poses a particular threat in that there is near loss of the intestinal epithelial barrier to contain this pathogen within this colonization niche. Several animal models have recapitulated these clinical circumstances by administration of chemotherapy and radiation doses accompanied by intestinal exposure to P. aeruginosa which results in death associated bacterial dissemination. In clinical studies, the P. aeruginosa blood isolates can be molecularly traced to isolates from feces, confirming the intestinal source of the bacteremia. In order to determine if a particular virulence determinant could be identified in blood isolates that predicted a particularly invasive phenotype against the intestinal epithelium, Yamaguchi et al. tested blood isolates of P. aeruginosa for their effect on human cultured intestinal epithelial cells. Results indicated that although several virulence determinants were identified, none appeared to confer a dissemination phenotype, although exotoxin A mutants were attenuated in this response.145 Again, it is important to recognize in studies in which severe immunocompromise exists and intestinal P. aeruginosa dissemination is present, that there are many cases where intestinal P. aeruginosa can cause systemic sepsis syndrome without being present in blood. Yet currently there is no epidemiologic evidence to point out each mechanism without using a molecular approach. Whether dissemination to remote organs is a surrogate marker for a high microbial burden within the intestine or itself a putative cause of systemic inflammation and organ damage is not known. Clinically it is impossible to dissociate the two phenomena and therefore this issue cannot be resolved. However antibiotics are usually administered intravenously in such cases, and sterilizing the bloodstream and systemic organs of P. aeruginosa does not necessarily eradicate the organism in the intestinal tract where it can exists in an impenetrable biofilm form. This observation may explain the continued high mortality rate when P. aeruginosa is isolated in the blood of patients following radiation, chemotherapy, or bone marrow transplantation despite the use of multiple antibiotic agents. Although the mechanisms by which intestinal P. aeruginosa causes mortality following radiation, chemotherapy, or bone marrow transplantation remain unknown, approaches to vaccinate patients are likely to fail given that protein synthesis is impaired at the time of exposure to the pathogen. Selective antibiotic decontamination of the gut prior

Molecular Detection of Foodborne Pathogens

to therapy is frought with antibiotic resistance outbreaks. Thus alternative therapies to combat these infections without focusing on pure microbiocidal effects may be warranted.

31.1.4 Novel Nonantibiotic Strategies to Contain Intestinal P. aeruginosa Since antibiotic treatment often fails to eradicate intestinal P. aeruginosa and can even lead to an opposite effect by triggering antibiotic resistance, novel approaches are needed to prevent lethal gut-derived sepsis due to this organism by focusing on the mechanisms that shift its phenotype during colonization from one of indolent resident to that of an adhesive, barrier dysregulating, disseminating strain. An integral part of this approach will involve enhancing detection of phenotype shifts by developing methods to molecularly type colonizing strains throughout the course of host vulnerability. In addition, approaches that are purely microbiocidal will have to be substituted for those designed to suppress virulence expression without eliminating the organism. Part of this approach will involve identifying the environmental cues, nutritional requirements, and host markers that shift P. aeruginosa virulence expression to identify the molecular signatures in the host and pathogen that signal a pathological interaction. Since there are likely multiple virulence determinants across several strains of P. aeruginosa that can cause intestinally derived sepsis and dissemination, common regulatory circuits of virulence expression will need to be identified that can be pharmacologically silenced during host stress. Since molecular detection techniques have identified P. aeruginosa to be present in the intestine of up to 20% of normal subjects, precisely how this pathogen shifts to an opportunistic phenotype within the gut has received little attention relative to its behavior in other sites such as the lung. There is a pressing need to preserve the normal intestinal flora during severe host stress as it has been shown to exert several probiotic and cytoprotective effects on the gut barrier and intestinal immune system. All antibiotics suffer from their collateral damage to the normal commensal microflora. A more strategic focus on how opportunistic pathogens shift their phenotypes to express harmfulness against the host could lead to strategies that interdict in the infectious process at its most proximate point.

31.1.5 Molecular Tools for P. aeruginosa Detection 31.1.5.1  P. aeruginosa Detection in Food Products Public concern regarding food safety has markedly increased over the past decade. Current Food Safety Inspection Service (FSIS) regulations for the food-processing industry include mandatory testing programs Salmonella, Listeria monocytogenes, Campylobacter, and Escherichia coli O157:H7 while low attention is paid today for P. aeruginosa. Science-based pathogen-specific tests can be divided into the categories of metabolic-based and genome-based methods.

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(i) Metabolic-based technologies: Direct enumeration of bacteria grown on agarized selective media. The principle is based on the direct enumeration of bacteria grown on agarized selective media. Different selective enrichment media, mainly Pseudomonas agar-based media were developed that allow the detection of P. aeruginosa in contaminated food samples. One of the examples of the specific modification of Pseudomonas agar-based media is that developed for detection of P. aeruginosa in mineral water that occasionally found to be contaminated in various countries such as Brazil, Canada, France, Germany, Spain, USA.146,147 Examination of drinking water for contamination with P. aeruginosa is not recommended as a routine procedure, but according to the European Directive for mineral water, natural mineral water should be free of P. aeruginosa in any 250-ml sample. Rita Ramalho et al.148 found that Pseudomonas agar-based media containing cetrimide, a specific selective agent for P. aeruginosa, is not suitable for detection of P. aeruginosa in mineral water. Cetrimide, a quaternary ammonium compound, that disrupts the bacterial cell membrane is generally not active against P. aeruginosa, however when in mineral water, cells become sensitive to this compound,148 possibly because of low nutrient effect on the cell wall permeability.149 Therefore, new media has been developed consisting of Pseudomonas Agar Base with added nalidixic acid, 15 mg/l (to suppress the growth of gram­positive bacteria) and 0.05% FeSO4 (to produce characteristic dark brown coloration in colonies in P. aeruginosa).148 The time required for the detection of P. aeruginosa by direct enumeration of bacteria is ~20 hrs but is not expansive and does not require any specific equipment. Direct impedance measurement. The direct impedance measurement is a modified strategy of metabolic-based methods that is based on the rapid detection of metabolic end-products of bacteria grown on specific media.150–153 This method is specifically useful for the rapid detection of P. aeruginosa in water, as a potential source of infection. The accumulation of metabolic end-products associates with corresponding changes in the resistance to the flow of a current (i.e., the impedance). The time required for the detection of impedance change is related to the initial concentration of bacteria in the water.154 The reported detection times ranged from 2–6 to 11–14 hrs.154,155 A specific highly selective media named as Z-broth has been developed for P. aeruginosa156 that relies on the ability of P. aeruginosa to grow on ammonia and acetate as the sole sources of nitrogen and carbon. Acetamide, the compound containing ammonia and acetate in an amide linkage, has been included as the main component of Z-broth that is dissociated by P. aeruginosa leading to the change in the impedance.156,157 Magnetoelastic biosensor for direct quantification of bacteria. Magnetoelasticity-based technology is yet an another example of metabolic-based methods that relies on the measurement of magnetic flux, a variance dependent on the bacterial density and changes in the composition of culture media during bacterial growth, in the response to an externally applied time-varying magnetic field. Magnetoelastic biosensor is a free standing, ribbon-like magnetoelastic thick-

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film coupled with a chemical or biochemical sensing thin film. In response to an externally applied time-varying magnetic field, the magnetoelastic ribbon mechanically vibrates at a characteristic frequency changing the magnetic flux that is detected specifically.158–160 Magnetoelastic biosensors have been developed for multiple chemical and biochemical analysis, such as the detection of ovidin,161 glucose,162 ammonia,163 pH,164 CO2,165, etc. A magnetoelastic immunosensor has been used for the direct, real-time quantification of P. aeruginosa in growth media.166 Changes in the properties of the culture medium due to consuming nutrients from a liquid culture medium and growth of bacteria lead to the changes in the resonance frequency of the sensor that are specifically measured. Fourier transform infrared spectroscopy (FT-IR) technology. FT-IR absorbance spectroscopy is an useful tool for early identification of growing bacteria.167–169 FT-IR spectroscopy measures vibrations of functional groups and highly polar bonds in DNA/RNA, proteins, cell wall amine- and fatty acid-containing components169 creating a “fingerprint” for a single bacterial strain. Rapid detection and identification of P. aeruginosa in bottled drinking water using using FT-IR spectroscopy and multivariate analysis has been reported.48 Electroimmunoassay technology. Electroimmunoassay technology is based on specific antibody-antigen binding to the production of an electrical signal. The technology is comprised of a circuit with a capture antibody attached to the solid surface. Upon addition of sample, the target antigen binds to the capture antibody. Commercially available is the Detex fully automated system (Molecular Circuitry; King of Prussia, PA) that consists of MC-18 immunoassay instrument and disposable pathogen-detection kits for various target pathogens (not yet for P. aeruginosa). As an immunoassay platform, the Detex system menu could be expanded to include tests for P. aeruginosa that are commercially available (Abcam Inc., Cambridge, MA; AbD Serotec, Oxford, UK; Affinity BioReagents, Golden, CO; Advanced Targeting Systems, San Diego, CA). (ii) Genotype-based methods: Culture-based identification and immunological assays use the phenotypic characteristics of the microorganism. However, identification criteria such as colony morphology, production of certain antigens (e.g., toxins), FT-IR fingerprint, impedance, magnitoelasticity could vary in between different isolates of P. aeruginosa, and on different media due to high variation of different P. aeruginosa isolates in the ability to produce pyocyanin, a blue color pigment with redox capabilities, pyoverdin, a fluorescence iron-scavenging peptide in P. aeruginosa, exotoxin A, a ADP-ribosyltransferase toxin, LPS, etc. These intrinsic (based on genome differences) and extrinsic (induced by conditions of cultivation) differences may lead to a misinterpretation of results and misidentification of bacteria. Genome-based identification methods using conserved regions of bacterial genomes use more stable genotypic characteristics of the microorganism.170 Besides, they are more sensitive allowing detecting single colonies without pre-cultivation. Nucleic-acid based identification of conserved regions or genes in the bacterial genome can

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be exploited for bacterial species identification while genes encoding specific virulence factors can be useful for defining the pathogenic potential of bacteria. Direct hybridization assay. Direct hybridization assay using labeled oligonucleotide probes currently are used for culture confirmation or for direct detection of organisms including P. aeruginosa in clinical or food samples.171–173 Basically, universal primers are used to amplify 16S–23S rDNA intergenic regions followed by hybridization of PCR product to species-specific oligonucleotides immobilized on a nylon membrane. Probe-based assays show a high degree of specificity because, when using stringent reaction conditions, a positive hybridization signal is correlated directly with the presence of the organism. A disadvantage of direct hybridization-based assays is a relatively large number of target cells requirement although using ribosomal ribonucleic acid (rRNA) as a target molecule can solve this problem. An oligonucleotide array using 16S–23S rDNA intergenic region has been successfully used for identification of P. aeruginosa.171 However, recent study of Lavenir et al.174 demonstrated that a more specific target gene can be used for P. aeruginosa detection in environmental samples. They have tested the reliability of the most widely used PCR screening for P. aeruginosa and found that 16S rDNA, oprI, oprL, and fliC were not specific enough to discriminate P. aeruginosa from 15 Pseudomonas species; gyrA, toxA, 16S–23S rDNA internal transcribed spacer (ITS) were demonstrated to be specific to discriminate against Pseudomonas species, however, toxA has not been found in 5% P. aeruginosa tested strains. Further efficacy screening gyrA (primers 5′-CCTGACCATCCGTCGCCACAAC-3′ and 5′-CGCAGCAGGATGCCGACGCC-3′) and 16S–23S rDNA ITS (primers 5′-TCCAAACAATCGTCGAAAGC-3′ and 5′-CCGAAAATTCGCGCTTGAAC-3′) in nine soils and 29 freshwater DNA extracts revealed uncertainties and false positive results. Therefore, they have developed a novel PCR screening, targeting the ecfX gene, encoding an ECF (extracytoplasmic function) sigma factor which is restricted to P. aeruginosa, and might play a role in heme-uptake and virulence. Specificity and sensitivity analyses have shown that ecfX (PA1300) PCR screening using primers 5′-ATGGATGAGCGCTTCCGTG-3′ and 5′-TCATCCTTCGCCTCCCTG-3′ is highly reliable, giving PCR products of the expected size for all P. aeruginosa strains tested and not amplifying DNA from any of the other Pseudomonas species tested, and was 100% reliable in soil and water DNA extracts.174 For more rapid control of bacterial food contamination, a new technique has been developed based on buoyant density gradient centrifugation to separate bacteria from complex food matrices, remove PCR inhibiting compounds, and concentrate contaminating food bacteria.175 Fluorescent in situ hybridization (FISH). FISH method uses fluorescent probes that bind specifically to a targeted DNA sequence that can be observed by fluorescence microscopy and flow cytometry. Traditionally, rRNA-­targeted probes are used, however, in most environmental samples FISH signal has a low signal-to-noise ratio because of low amount of bacteria.

Molecular Detection of Foodborne Pathogens

A recent advantage in FISH technology is the replacement of standard linear oligonucleotide probes with molecular beacons. The use of DNA molecular beacons for flow cytometry detection of P. putida has doubled signal-to-noise ratio demonstrating improved detection in spiked environmental samples, without a need for separate washing steps, and have allowed detection in activated sludge and river water samples.176 Real-time fluorescent PCR technologies. Amplification technologies combined with fluorescent-labeled probes have strongly contributed to the acceptance of molecular assays in the microbiology laboratories because of high specificity and sensitivity of this method. Most of these technologies are based on the principle of fluorescence resonance energy transfer and require real-time thermocycling instruments. This technology known as the TaqMan® system uses 5´ to 3´ exonuclease activity of Taq DNA polymerase. An oligonucleotide probe is labeled with a fluorescent reporter dye at the 5´ end and a fluorescent quencher dye at the 3´ end. Upon hybridization of the probe molecule to the target DNA during amplification, the 5´ nuclease activity of the enzyme cleaves the probe, separating the reporter dye from the quencher dye that results in increase of fluorescence. PCR thermal cycling exponentially amplifies target DNA that is reflected by fluorescence intensity. Using TaqMan® real-time quantitative PCR, P. aeruginosa has been successfully detected during examination of municipal wastewater treatment together with another 13 foodborne pathogens.177 The primers and TaqMan® probe were designed for regA gene (GeneBank accession no. X123666) of P. aeruginosa: F primer 5′-TGCTGGTGGCACAGGACAT-3′, R primer 5′-TTGTTGGTGCAGTTCCTCATTG-3′, and TaqMan® probe 5′-CAGATGCTTTGCCTCAA-3′.177 The following technique of bacterial DNA isolation has been used by authors: DNA was extracted from biomass collected from 200 ml of raw wastewater pre-treated with 120 μl of a 10 mg/ml lysozyme-50 mM EDTA solution, followed by the addition of 480 μl of 50 mM EDTA. The samples were incubated in a water bath at 37°C for 45 min, and centrifuged at 16,000 × g for 2 min, and the supernatant removed. Genomic DNA was extracted from pre-treated wastewater samples and pure culture samples using the Wizard® Genomic DNA Purification Kit (Promega™ Corp., Madison, WI). A ribonuclease A digestion is included in this extraction protocol. Due to the large amounts of impurities contained in the wastewater DNA samples, the protocol was repeated again from the protein precipitation step, to the ethanol purification step to further purify DNA. The quantity and purity of the DNA extract was determined spectrophotometrically at an absorbance of 260 nm, and A260/A280, respectively. Before pure culture or wastewater genomic DNA was used for real-time PCR, it was further purified using the Qiagen® Genomic-Tip 20/G Purification Kit (QIAGEN® Inc. Canada, Mississauga, ON, Canada), which was carried out according the manufacturer’s protocol. The quantity and purity of the DNA extract was measured using a spectrophotometer as described previously. Template genomic DNA, primers and TaqMan® probes were added to TaqMan® Universal PCR Mastermix (Applied Biosystems, Foster City, CA) to a final concentration of 250

Pseudomonas

nM TaqMan® probes, and 1 μM for each primer. Real-time PCR reactions were executed in a SmartCycler® II System (Cepheid Inc., Sunnyvale, CA) with a temperature profile of 50°C for 2 min, 95°C for 10 min, followed by 45 cycles at 95°C for 15 sec and 60°C for 60 sec.177 31.1.5.2 Detection of P. aeruginosa in Hospitals Clinical Laboratories The current methods for P. aeruginosa identification in hospitals clinical laboratories are mainly metabolic-based techniques that identify bacteria based on biochemical tests enabling the identification of a wide variety of different bacteria and antibiotic susceptibility testing.178,179 The world-widely used automated VITEK 2 system (bioMérieux) includes a software system that allows identification and antibiotic resistance patterns of bacteria. Although VITEK 2 system allows fast identification process of ~3–17 hrs, the requirement to use pure bacterial culture for identification with this system prolongs the whole procedure of analysis of clinical samples up to 2 days. Besides, VITEC 2 system is useless for identification of antibiotic resistance in mucoid P. aeruginosa strains. In this case, the susceptibility has to be determined by the disc diffusion method on Mueller–Hinton II agar plates (Becton Dickinson) using antibiotic-containing discs. Overall, minimum time requirement for clinical sample analysis to identify bacteria and antibiotic resistance profile from the point of clinical sample arrival is 36–48 hrs for nonmucoid and 48–60 hrs for mucoid strains of P. aeruginosa. To identify bacteria directly in blood cultures and body fluids, BacT/Alert Microbial Detection System (BacT/ALERT 3D) is used. BacT/Alert is an automated microbial detection system based on the detection of CO2 produced by growing microorganisms.180,181 It has high level sensitivity allowing detection of single bacterial cells in 1 ml blood, however dependently on the titer and growth rate, it might take from hours to up to 5 days to conclude bacteremia. In the case of positive test, bacteria is subjected to Gram test following by plating on the specific agarized plates, incubating overnight and then followed by VITEK 2 identification and antibiotic resistance. Above described procedures common in hospital clinical laboratories are reliable and allow the identification of the infecting bacteria and antibiotic for its treatment. However, such analyses are not able to provide “an express” bacterial identification and antibiotic susceptibility sometimes required to be done during few hours. In this case, molecular techniques that have been discussed above are necessary to apply. The first important step in molecular-based diagnostics is the collection, handling, and storage of intestinal samples that can include feces and biopsies. The most detailed protocols for collection and handling of intestinal samples as well as DNA isolation are described in Nature Protocols by Zoetendal et al.182 The protocols developed for gastrointestinal tract samples including feces, biopsies, and ileostomy effluents are based on mechanical disruption, followed by ­isolation of nucleic acids using phenol:chloroform:isoamy­ lalcohol extraction, or use of an alternative DNA isolation protocol that is based on a commercial kit (Qiagen). Timing

439

of 2.5–4 hrs is required for total procedure. Another alternative rapid extraction method, Bead Beating Plus CHELEX (BB + C), is avoided of using phenol:chloroform:isoamy­ lalcohol extraction, however it was not tested with feces samples.183 Multiple methods for fecal preservation and DNA extraction were tested by Nechvatal et al.184 They paid specific attention to the presence of PCR inhibitors in fecal extracts185 that include bilirubins, bile salts, and complex carbohydrates.186 At least 100-fold dilution was needed to successfully amplify DNA from extracts.184 Most of molecular approaches use 16S rRNA as a phylogenetic marker for the identification of microbes because 16S rRNA presents in every cell and the structure of its gene is conserved since has a low mutation and horizontal transfer rate. Multiple reports demonstrated the usefulness of 16S rRNA-based specific identification of P. aeruginosa clinical isolates: (i) partial 16S rRNA gene sequencing to verify the identification of clinical isolates that could not be clearly determined with routine tests;187 (ii) PCR amplification of a 233 bp fragment in 16S rRNA gene (with primers 5′-GAG GAA GGT GGG GAT GAC GT-3′ and 5′-AGG CCC GGG AAC GTA TTC AC-3′) to allow cost effective and fast identification of P. aeruginosa in complex clinical samples. However, based on the results of Lavenir et al. study, ecfX gene might represent a better target gene for P. aeruginosa identification that should be verified using clinical samples. 31.1.5.3 Assessment of Virulence Potential in P. aeruginosa P. aeruginosa is often difficult to eliminate by antibiotics because of its intrinsic antibiotic resistance that rapidly can be transformed to multi-antibiotic resistance, and because of the ability of this organism to form biofilm under stressed conditions. A biofilm is a complex aggregation of microorganisms representing cell communities that characterized by surface attachment, structural heterogeneity, genetic diversity, complex community interactions, and an extracellular matrix of polymeric substances that drastically contributes to antibiotic resistance.188,189 Thus, treatment additions to antibiotics are needed based on a detailed analysis of the virulence potential and specific gene array expression profile. The virulence potential could be mainly judged by the presence of pathogenic gene island in P. aeruginosa containing exoU gene. About 30% of environmental and clinical P. aeruginosa strains harbor the exoU gene that appears to significantly correlate with a cytotoxic and lethal phenotype. The exoU gene encodes the most toxic effector protein of type III secretion system that is injected directly through a needle-like system in the cytoplasm of host eukaryotic cells. The presence of exoU in P. aeruginosa can be easily detected by including specific primer set exoU2998, 5´-GCTAAGGCTTGGCGGAATA-3´ and exoU3182, 5´-AGATCACACCCAGCGGTAAC-3´. Pre­ventive techniques to attenuate the adhesiveness of P. aeruginosa to intestinal epithelial cells have the potential to decrease the risk of mortality due to P. aeruginosa strains harboring exoU gene. Analysis of the transcriptional activation of specific virulence subsystems might allow for novel

440

strategies to down-regulate these systems as they become in vivo activated. The most detailed protocol for RNA isolation from bacterial cells of the human gastrointestinal tract is described in Nature Protocols by Zoetendal et al.190

31.2  Methods 31.2.1 Sample Preparation Bacterial DNA isolation from culture (1) Grow culture of P. aeruginosa on liquid TSB media to OD = 0.6. (2) Remove 5 ml of culture to a tube, centrifuge at 5000 rpm for 5 min, discard supernatant, and purify genomic DNA as described for fecal samples (from step 2). Bacterial DNA isolation from human feces Prior DNA extraction, samples could be lyophilized. (1) Weight 200 µg of fresh or 30 µg of lyophilized feces. (2) Deposit sample into 2 ml Eppendof tube containing three to five glass beads (3 mm). (3) Add 800 µl TNES buffer and 10 µl 20 mg/ml proteinase K (Invitrogen, Cat No. 25530-049) to the stool sample, vigorously shake for 3 min (TNES buffer: 50 mM Tris-HCl pH 7.4, 100 mM EDTA pH 8.0, 400 mM NaCl, 0.5% SDS). (4) Incubate overnight at 55°C. (5) Add 200 µl 6 M NaCl and shake by vortex for 30 sec. (6) Centrifuge at maximum speed for 1 min. Transfer supernatant into the new 1.5 ml Eppendorf tube. At this point, samples can be frozen and stored at –20°C. (7) Take 500 µl of supernatant from step #6, add 500 µl of phenol-chlorophorm:isoamylalcohol (25:24:1) (Invitrogen, Cat No.15593-031), mix by vortexing, and centrifuge at 12,000 rpm for 5 min. Transfer the aqueous phase to the new Eppendorf tube. (8) Add 500 µl 100% ethanol to the tube containing aqueous phase from step 8 and mix by vortexing. (9) Centrifuge 12,000 rpm, 4°C for 10 min. Decant supernatant. (10) Add 70% ethanol, mix by vortexing and centrifuge at 12,000 rpm, 4°C, 10 min. Decant supernatant and carefully remove the residual liquid by pipetting. (11) Air dry at room temperature. (12) Add 100 µl of sterile water or TE buffer to dissolve DNA. (TE buffer: 10 mM Tris-HCl pH 7.6, 1 mM EDTA pH 8.0). (13) Adjust concentration to 50 ng/µl. Store –80°C.

31.2.2  Detection Procedure We present below a real-time quantitative PCR (qPCR) assay with primers (5′-CAAAACTACTGAGCTAGAGTACG-3′

Molecular Detection of Foodborne Pathogens

and 5′-TAAGATCTCAAGGATCCCAACGGCT-3′) targeting 23S rRNA gene of P. aeruginosa that are based on the report of Matsuda et al.191 (1) Prepare PCR mixture (10 µl) containing 10 µl of SYBR Green PCR mixture (Part No. 4309155, Applied Biosystems, Foster City, CA), 1 µl of each primer (10 µM), 1 µl of DNA, and 7 µl of water. (2) Run PCR in ABI PRISM 7900HT sequence detection system (Applied Biosystems) with the follow cycling: one cycle of 95°C for 15 min; 40 cycles of 94°C for 20 sec, 60°C for 20 sec, 72°C for 50 sec; one cycle of 94°C for 15 sec. (3) Detect the fluorescent products in the last step of each cycle. Perform a melting curve analysis after amplification to distinguish the target from the nontargeted PCR product, by slow heating at temperatures from 60 to 95°C at a rate of 0.2°C/sec with continuous fluorescence collection.

31.3  Conclusion Multiple lines of evidence continue to accumulate that demonstrate that in a susceptible host, intestinal P. aeruginosa is capable of causing a wide spectrum of infection related diseases including diarrhea, intestinal necrosis, dissemination to remote organs, and severe sepsis and shock. In rare cases, even in a normal host, P. aeruginosa has been reported to cause fatal sepsis from within the intestinal tract. Since as many as 20% of normal subject are intestinally colonized with this pathogen, a better understanding of the molecular determinants that shift this bacterium from indolent colonizer to disease producing pathogen will be necessary to develop detection markers and therapeutic strategies to contain its pathogenicity. The persistent problem of MDR begs alternative approaches that can prevent and treat P. aeruginosa without the use of antibiotics. Such approaches will require a more complete understanding of the molecular mechanisms and environmental cues by which P. aeruginosa shifts its phenotype during the course of human stress and illness when present in the intestine.

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443 102. Tascini, C. et al. Microbiological activity and clinical efficacy of a colistin and rifampin combination in multidrug-resistant Pseudomonas aeruginosa infections. J. Chemother., 16, 282, 2004. 103. Ferrara, A.M. Potentially multidrug-resistant non-fermentative Gram-negative pathogens causing nosocomial pneumonia. Int. J. Antimicrob. Agents, 27, 183, 2006. 104. Zaborina, O. et al. Identification of multi-drug resistant Pseudomonas aeruginosa clinical isolates that are highly disruptive to the intestinal epithelial barrier. Ann. Clin. Microbiol. Antimicrob., 5, 14, 2006. 105. Khakoo, R.A. and Kluge, R.M. Decreased virulence of gentamicin-resistant strains of Pseudomonas aeruginosa in a rat model. J. Lab. Clin. Med., 91, 96, 1978. 106. Keys, T. F. and Washington, J. A., 2nd. Gentamicin-resistant Pseudomonas aeruginosa: Mayo Clinic experience, 1970– 1976. Mayo Clin. Proc., 52, 797, 1977. 107. Hostacka, A. et al. Clinical pseudomonas aeruginosa: potential factors of pathogenicity and resistance to antimicrobials. Folia Microbiol. (Praha), 51, 633, 2006. 108. Linares, J.F. et al. Overexpression of the multidrug efflux pumps MexCD-OprJ and MexEF-OprN is associated with a reduction of type III secretion in Pseudomonas aeruginosa. J. Bacteriol. 187, 1384, 2005. 109. Hocquet, D. et al. Pseudomonas aeruginosa may accumulate drug resistance mechanisms without losing its ability to cause bloodstream infections. Antimicrob. Agents Chemother., 51, 3531, 2007. 110. Ruiz, J. Mechanisms of resistance to quinolones: target alterations, decreased accumulation and DNA gyrase protection. J. Antimicrob. Chemother., 51, 1109, 2003. 111. Jeannot, K. et al. Induction of the MexXY efflux pump in Pseudomonas aeruginosa is dependent on drug-ribosome interaction. J. Bacteriol., 187, 5341, 2005. 112. Li, X.Z., Poole, K. and Nikaido, H. Contributions of MexAB-OprM and an EmrE homolog to intrinsic resistance of Pseudomonas aeruginosa to aminoglycosides and dyes. Antimicrob. Agents Chemother., 47, 27, 2003. 113. Hauser, A.R. and Sriram, P. Severe Pseudomonas aeruginosa infections: tackling the conundrum of drug resistance. Postgrad. Med., 117, 41, 2005. 114. Pearson, J.P., Van Delden, C. and Iglewski, B.H. Active efflux and diffusion are involved in transport of Pseudomonas aeruginosa cell-to-cell signals. J. Bacteriol. 181, 1203, 1999. 115. Kohler, T. et al. Overexpression of the MexEF-OprN multidrug efflux system affects cell-to-cell signaling in Pseudomonas aeruginosa. J. Bacteriol.,183, 5213, 2001. 116. Stavrinides, J., McCann, H.C. and Guttman, D.S. Hostpathogen interplay and the evolution of bacterial effectors. Cell. Microbiol., 10, 285, 2008. 117. Mota, L.J. and Cornelis, G.R. The bacterial injection kit: type III secretion systems. Ann. Med., 37, 234, 2005. 118. Kubori, T. et al. Molecular characterization and assembly of the needle complex of the Salmonella typhimurium type III protein secretion system. Proc. Natl. Acad. Sci. USA, 97, 10225, 2000. 119. Hueck, C.J. Type III protein secretion systems in bacterial pathogens of animals and plants. Microbiol. Mol. Biol. Rev., 62, 379, 1998. 120. Sanchez, P. et al. Fitness of in vitro selected Pseudomonas aeruginosa nalB and nfxB multidrug resistant mutants. J. Antimicrob. Chemother., 50, 657, 2002.

444 121. Cosson, P. et al. Pseudomonas aeruginosa virulence analyzed in a Dictyostelium discoideum host system. J. Bacteriol. 184, 3027, 2002. 122. Zaborina, O. et al. Structure-function aspects of PstS in multidrug-resistant Pseudomonas aeruginosa. PLoS Pathog., 4, e43, 2008. 123. Alverdy, J. et al. Gut-derived sepsis occurs when the right pathogen with the right virulence genes meets the right host: evidence for in vivo virulence expression in Pseudomonas aeruginosa. Ann. Surg., 232, 480, 2000. 124. Laughlin, R.S. et al. The key role of Pseudomonas aeruginosa PA-I lectin on experimental gut-derived sepsis. Ann. Surg., 232, 133, 2000. 125. Datta, H.K., Malik, M. and Neely, R.D. Hepatic surgeryrelated hypophosphatemia. Clin. Chim. Acta, 380, 13, 2007. 126. Martinez, M. J. et al. Hypophosphatemia in postoperative patients with total parenteral nutrition: influence of nutritional support teams. Nutr Hosp 21, 657-60 (2006). 127. Miller, D.W. and Slovis, C.M. Hypophosphatemia in the emergency department therapeutics. Am. J. Emerg. Med., 18, 457, 2000. 128. Tlaskalova-Hogenova, H. et al. Commensal bacteria (normal microflora), mucosal immunity and chronic inflammatory and autoimmune diseases. Immunol. Lett., 93, 97, 2004. 129. Wu, L. et al. Pseudomonas aeruginosa expresses a lethal virulence determinant, the PA-I lectin/adhesin, in the intestinal tract of a stressed host: the role of epithelia cell contact and molecules of the quorum sensing signaling system. Ann. Surg., 238, 754, 2003. 130. Fang, X.Q., Wang, W.F. and Liu, Y.N. [The role of the quorum sensing system in a rat model of Pseudomonas aeruginosa pulmonary infection and its relationship with the expression of virulence factors]. Zhonghua Jie He He Hu Xi Za Zhi, 29, 458, 2006. 131. Plotkowski, M.C. et al. Early mitochondrial dysfunction, superoxide anion production, and DNA degradation are associated with non-apoptotic death of human airway epithelial cells induced by Pseudomonas aeruginosa exotoxin A. Am. J. Respir. Cell. Mol. Biol., 26, 617, 2002. 132. Schultz, M.J. et al. Impairment of host defence by exotoxin A in Pseudomonas aeruginosa pneumonia in mice. J. Med. Microbiol., 50, 822, 2001. 133. Hirakata, Y. et al. Effect of Pseudomonas aeruginosa exotoxin A on endotoxin-induced tumour necrosis factor production in murine lung. J. Med. Microbiol., 48, 471, 1999. 134. Cummings, J.H. and Macfarlane, G.T. The control and consequences of bacterial fermentation in the human colon. J. Appl. Bacteriol., 70, 443, 1991. 135. Gibson, G.R. and Roberfroid, M.B. Dietary modulation of the human colonic microbiota: introducing the concept of prebiotics. J. Nutr. 125, 1401, 1995. 136. Donskey, C.J. The role of the intestinal tract as a reservoir and source for transmission of nosocomial pathogens. Clin. Infect. Dis., 39, 219, 2004. 137. Edlund, C. and Nord, C.E. Effect on the human normal microflora of oral antibiotics for treatment of urinary tract infections. J. Antimicrob. Chemother., 46 (Suppl. 1), 41; Discussion, 63, 2000. 138. Wu, L.R. et al. Surgical injury and metabolic stress enhance the virulence of the human opportunistic pathogen Pseudomonas aeruginosa. Surg. Infect. (Larchmt), 6, 185, 2005. 139. Patel, N.J. et al. Recognition of intestinal epithelial HIF1alpha activation by Pseudomonas aeruginosa. Am. J. Physiol. Gastrointest. Liver Physiol. 292, G134, 2007.

Molecular Detection of Foodborne Pathogens 140. Alverdy, J., Chi, H.S. and Sheldon, G.F. The effect of parenteral nutrition on gastrointestinal immunity: the importance of enteral stimulation. Ann. Surg. 202, 681, 1985. 141. Alverdy, J., Zaborina, O. and Wu, L. The impact of stress and nutrition on bacterial-host interactions at the intestinal epithelial surface. Curr. Opin. Clin. Nutr. Metab. Care, 8, 205, 2005. 142. Zaborina, O. et al. Dynorphin activates quorum sensing quinolone signaling in Pseudomonas aeruginosa. PLoS Pathog., 3, e35, 2007. 143. Wu, L. et al. Recognition of host immune activation by Pseudomonas aeruginosa. Science, 309, 774, 2005. 144. Kohler, J. E. et al. Components of intestinal epithelial hypoxia activate the virulence circuitry of Pseudomonas. Am. J. Physiol. Gastrointest. Liver Physiol., 288, G1048, 2005. 145. Hirakata, Y. et al. Adherence to and penetration of human intestinal Caco-2 epithelial cell monolayers by Pseudomonas aeruginosa. Infect. Immun., 66, 1748, 1998. 146. Hunter, P.R. The microbiology of bottled natural mineral waters. J. Appl. Bacteriol.,74, 345, 1993. 147. Warburton, D.W., Bowen, B. and Konkle, A. The survival and recovery of Pseudomonas aeruginosa and its effect upon salmonellae in water: methodology to test bottled water in Canada. Can. J. Microbiol., 40, 987, 1994. 148. Ramalho, R. et al. Modified Pseudomonas agar: new differential medium for the detection/enumeration of Pseudomonas aeruginosa in mineral water. J. Microbiol. Methods, 49, 69, 2002. 149. Goto, S. and Enomoto, S. Nalidixic acid cetrimide agar. A new selective plating medium for the selective isolation of Pseudomonas aeruginosa. Jpn J. Microbiol., 14, 65, 1970. 150. Colquhoun, K.O., Timms, S. and Fricker, C.R. Detection of Escherichia coli in potable water using direct impedance technology. J. Appl. Bacteriol., 79, 635, 1995. 151. Chang, T.C. and Huang, A.H. Rapid differentiation of fermentative from nonfermentative gram-negative bacilli in positive blood cultures by an impedance method. J. Clin. Microbiol., 38, 3589, 2000. 152. Wawerla, M. et al. Impedance microbiology: applications in food hygiene. J. Food Prot., 62, 1488, 1999. 153. Flint, S.H. and Brooks, J.D. Rapid detection of Bacillus stearothermophilus using impedance-splitting. J. Microbiol. Methods, 44, 205, 2001. 154. Cady, P. et al. Impedimetric screening for bacteriuria. J. Clin. Microbiol., 7, 273, 1978. 155. Silverman, M.P. and Munoz, E.F. Automated electrical impedance technique for rapid enumeration of fecal coliforms in effluents from sewage treatment plants. Appl. Environ. Microbiol., 37, 521, 1979. 156. Szita, G. et al. Detection of Pseudomonas aeruginosa in water samples using a novel synthetic medium and impedimetric technology. Lett. Appl. Microbiol.,.45, 42, 2007. 157. Szita, G. et al. A novel, selective synthetic acetamide containing culture medium for isolating Pseudomonas aeruginosa from milk. Int. J. Food Microbiol., 43, 123, 1998. 158. Schmidt, S. and Grimes, C.A. Characterization of nanodimensional thin-film elastic moduli using magnetoelastic sensors. Sens. Actuators A Phys., 94, 189, 2001. 159. Zourob, M. et al. A wireless magnetoelastic biosensor for the direct detection of organophosphorus pesticides. Analyst, 132, 338, 2007. 160. Ong, K.G. et al. A rapid highly-sensitive endotoxin detection system. Biosens. Bioelectron., 21, 2270, 2006. 161. Ruan, C. et al. A magnetoelastic bioaffinity-based sensor for avidin. Biosens. Bioelectron., 19, 1695, 2004.

Pseudomonas 162. Cai, Q. et al. A wireless, remote query glucose biosensor based on a pH-sensitive polymer. Anal. Chem., 76, 4038, 2004. 163. Cai, Q.Y., Jain, M. K. and Grimes, C.A. A wireless, remote query ammonia sensor. Sens. Actuators B Chem., 77, 614, 2001. 164. Cai, Q.Y. and Grimes, C.A. A remote query magnetoelastic pH sensor. Sens. Actuators B Chem., 71, 112, 2000. 165. Cai, Q.Y., Cammers-Goodwin, A. and Grimes, C.A. A wireless, remote query magnetoelastic CO2 sensor. J. Environ. Monit., 2, 556, 2000. 166. Pang, P. et al. Detection of Pseudomonas aeruginosa using a wireless magnetoelastic sensing device. Biosens. Bioelectron., 23, 295, 2007. 167. Sandt, C. et al. FT-IR microspectroscopy for early identification of some clinically relevant pathogens. J. Appl. Microbiol., 101, 785, 2006. 168. Al-Qadiri, H.M. et al. Fourier transform infrared spectroscopy, detection and identification of Escherichia coli O157:H7 and Alicyclobacillus strains in apple juice. Int. J. Food Microbiol., 111, 73, 2006. 169. Wenning, M., Theilmann, V. and Scherer, S. Rapid analysis of two food-borne microbial communities at the species level by Fourier-transform infrared microspectroscopy. Environ. Microbiol., 8, 848, 2006. 170. Jannes, G. and De Vos, D. A review of current and future molecular diagnostic tests for use in the microbiology laboratory. Methods Mol. Biol.,345, 1, 2006. 171. Lin, M.C. et al. Use of oligonucleotide array for identification of six foodborne pathogens and Pseudomonas aeruginosa grown on selective media. J. Food Prot., 68, 2278, 2005. 172. Wang, X.W. et al. Development and application of an oligonucleotide microarray for the detection of food-borne bacterial pathogens. Appl. Microbiol. Biotechnol., 76, 225, 2007. 173. Hong, B.X. et al. Application of oligonucleotide array technology for the rapid detection of pathogenic bacteria of foodborne infections. J. Microbiol. Methods, 58, 403, 2004. 174. Lavenir, R. et al. Improved reliability of Pseudomonas aeruginosa PCR detection by the use of the species-specific ecfX gene target. J. Microbiol. Methods, 70, 20, 2007. 175. Fukushima, H. et al. Rapid separation and concentration of food-borne pathogens in food samples prior to quantification by viable-cell counting and real-time PCR. Appl. Environ. Microbiol., 73, 92, 2007. 176. Lenaerts, J., Lappin-Scott, H.M. and Porter, J. Improved fluorescent in situ hybridization method for detection of bacteria from activated sludge and river water by using DNA molecular beacons and flow cytometry. Appl. Environ. Microbiol., 73, 2020, 2007.

445 177. Shannon, K.E. et al. Application of real-time quantitative PCR for the detection of selected bacterial pathogens during municipal wastewater treatment. Sci. Total Environ., 382, 121, 2007. 178. Joyanes, P. et al. Evaluation of the VITEK 2 system for the identification and susceptibility testing of three species of nonfermenting gram-negative rods frequently isolated from clinical samples. J. Clin. Microbiol., 39, 3247, 2001. 179. Ling, T.K. et al. Evaluation of VITEK 2 rapid identification and susceptibility testing system against gram-negative clinical isolates. J. Clin. Microbiol., 39, 2964, 2001. 180. Thorpe, T.C. et al. BacT/Alert: an automated colorimetric microbial detection system. J. Clin. Microbiol., 28, 1608, 1990. 181. Chryssanthou, E., Cars, O. and Sjolin, J. New automated method for determining postantifungal effect of amphotericin B against Candida species: effects of concentration, exposure time, and area under the curve. Antimicrob. Agents Chemother., 46, 4016, 2002. 182. Zoetendal, E.G. et al. Isolation of DNA from bacterial samples of the human gastrointestinal tract. Nat. Protoc., 1, 870, 2006. 183. Jaffe, R.I., Lane, J.D. and Bates, C.W. Real-time identification of Pseudomonas aeruginosa direct from clinical samples using a rapid extraction method and polymerase chain reaction (PCR). J. Clin. Lab. Anal., 15, 131, 2001. 184. Nechvatal, J.M. et al. Fecal collection, ambient preservation, and DNA extraction for PCR amplification of bacterial and human markers from human feces. J. Microbiol. Methods, 72, 124, 2008. 185. Flekna, G. et al. Real-time PCR method with statistical analysis to compare the potential of DNA isolation methods to remove PCR inhibitors from samples for diagnostic PCR, Mol. Cell. Probes, 21, 282, 2007. 186. Monteiro, L. et al. Complex polysaccharides as PCR inhibitors in feces: Helicobacter pylori model, J. Clin. Microbiol., 35, 995, 1997. 187. Moissenet, D. et al. [Use of 16S rRNA gene sequencing for identification of “Pseudomonas-like” isolates from sputum of patients with cystic fibrosis], Pathol. Biol. (Paris), 53, 500, 2005. 188. Mah, T.F. and O’Toole, G.A. Mechanisms of biofilm resistance to antimicrobial agents, Trends Microbiol., 9, 34, 2001. 189. Mah, T.F. et al. A genetic basis for Pseudomonas aeruginosa biofilm antibiotic resistance, Nature, 426, 306, 2003. 190. Zoetendal, E.G. et al. Isolation of RNA from bacterial samples of the human gastrointestinal tract. Nat. Protoc., 1, 954, 2006. 191. Matsuda, K. et al. Sensitive quantitative detection of commensal bacteria by rRNA-targeted reverse transcription-PCR. Appl. Environ. Microbiol., 73, 32, 2007.

32 Salmonella

Charlotta Löfström, Jeffrey Hoorfar Technical University of Denmark

Jenny Schelin, Peter Rådström Lund University

Burkhard Malorny

Federal Institute for Risk Assessment

Contents 32.1 Introduction.................................................................................................................................................................... 447 32.1.1 Biology.............................................................................................................................................................. 447 32.1.2 Pathogenesis...................................................................................................................................................... 448 32.1.3 Epidemiology.................................................................................................................................................... 448 32.1.4 Conventional Diagnostics................................................................................................................................. 449 32.1.5 Molecular Diagnostics...................................................................................................................................... 450 32.1.5.1 Immunological Methods.................................................................................................................. 451 32.1.5.2 Nucleic Acid-Based Methods........................................................................................................... 452 32.2 Methods.......................................................................................................................................................................... 453 32.2.1 Sample Collection and Preparation.................................................................................................................. 453 32.2.1.1 Sample Collection............................................................................................................................ 453 32.2.1.2 Sample Preparation.......................................................................................................................... 453 32.2.2 Detection Procedures........................................................................................................................................ 454 32.3 Conclusions and Future Perspectives............................................................................................................................. 455 Acknowledgments...................................................................................................................................................................... 456 References.................................................................................................................................................................................. 456

32.1  Introduction 32.1.1  Biology Salmonellae are rod-shaped facultative anaerobic, catalasepositive, oxidase-negative, Gram-negative bacteria which are usually motile. The bacteria are nonspore forming with a size of 0.7–1.5 μm × 2.0–5.0 mm producing colonies generally 2–4 mm in diameter.1 The genus Salmonella is a member of the family Enterobacteriaceae and is named after the American bacteriologist D.E. Salmon who identified S. choleraesuis in 1885.1 Two species are currently recognized, namely Salmonella enterica and Salmonella bongori (formerly subspecies V). S. enterica has been subdivided into six subspecies, S. enterica subsp. enterica (designated subspecies I), S. enterica subsp. salamae (subspecies II), S. enterica subsp. arizonae (subspecies IIIa), S. enterica subsp. diarizonae (subspecies IIIb), S. enterica subsp. houtenae (subspecies IV) and S. enterica subsp. indica (subspecies VI).1,2 A number of biochemical reactions are used to differentiate between the species and subspecies.3 Subspecies I strains are

usually isolated from humans and warm-blooded animals and the other subspecies usually from cold-blooded animals and the environment. According to the Kauffmann–White scheme,4,5 subspecies are further divided into serotypes which are widely used as an epidemiological standardized typing method. Serotyping is based on the antigenic variability at lipopolysaccharide moieties (O antigens), flagellar proteins (H1 and H2 antigens), and capsular polysaccharides (Vi antigens). Currently over 2500 different serovars are known by this scheme. Salmonella serovars may be found predominantly in one host, may be ubiquitous or may have an unknown host. Salmonella can also be found in the natural environment, e.g., water, soil, and plants, but does not seem to grow significantly. The bacteria can, however, survive several weeks in water and several years in soil if the temperature, humidity, and pH are favorable.6 Salmonella serovars may be hostspecific, host-restricted or ubiquitous. Thus, serovar Typhi, serovar Gallinarum, and serovar Abortusovis are almost exclusively associated with systemic disease in humans, 447

448

fowl, and sheep, respectively. Host-restricted strains, while primarily associated with one or two closely related host species, may also infrequently cause disease in other hosts. For example, serovar Dublin and serovar Choleraesuis are generally associated with severe systemic disease in ruminants and pigs, respectively. Ubiquitous serovars, such as serovar Typhimurium and serovar Enteritidis, usually induce gastroenteritis in a broad range of unrelated host species.

32.1.2  Pathogenesis In general, S. enterica from human infections can be subdivided into two groups: the enteric fever (typhoidal) group and the nontyphoidal salmonellae, which typically cause gastroenteritis. Symptoms appear within 6–24 h after ingestion of contaminated food or water and last for as long as one week. Initial symptoms include nausea and vomiting, followed by abdominal pain and diarrhea and sometimes fever. Occasionally salmonellae can cause invasive (systemic) disease under certain conditions. The systemic disease can be fatal if untreated and kills about 10% of infected people. The incubation period is longer than for nontyphoidal salmonellae (approximately 1 week). Invasive salmonellae enter the blood stream and spread throughout the body. The symptoms are high fever (typhoid fever) and anorexia caused by LPS-mediated release of cytokines. Ills, convulsions, and delirium can also appear. Mainly, five serotypes are involved in typhoid fever, namely Typhi, Paratyphi A, Paratyphi B, Paratyphi C, and related serovars (Choleraesuis and Sendai).7 The other approximately 2500 known serotypes belong to the nontyphoidal salmonellae. Although rare, nontyphoidal salmonellae can cause systemic disease, typically when the host’s defense is compromised. Specific nontyphoidal serotypes appear to be associated with rather high ratios of invasiveness compared to other serotypes, e.g., Dublin, Heidelberg, Brandenburg, and Virchow.8 Clearly the nature and severity of Salmonella infections in different animals and humans vary enormously and are influenced by many factors, including the infecting Salmonella serovar, strain virulence, infecting dose, host animal species, age, and immune status of the host, and geographical region. The route of salmonellae during infection can be manifold and despite enormous research activities the pathogenicity of Salmonella is still not well enough known. Usually, salmonellae colonize the intestine by the adhesion of the bacteria to the epithelial cells using fimbrial antigens. The cells invade the intestinal mucosa and multiply in the gut-associated lymphoid tissue (GALT). From the infected tissues the pathogens spread to the regional lymph nodes, where macrophages form a first effective barrier to prevent a further spread. If the marcrophages are unable to avoid the spread, systemic disease can occur. During the systemic disease the bacteria spread from the GALT via the efferent lymphatics and the thoracic duct into the vena cava. From there the salmonellae spread throughout the body. The bacteria multiply in spleen and liver and are released in large numbers to the blood stream infecting other organs. Salmonella are able to survive and to multiply inside host cells.

Molecular Detection of Foodborne Pathogens

Salmonella harbors large clusters of virulence genes that act together in a complex virulence function for different outcomes of Salmonella infections. They have been chromosomally acquired by horizontal gene transfer and are called pathogenicity islands. These islands contain genes required for the different roles in gastrointestinal and systemic pathogenesis.9 Some of them encode type III secretion systems (TTSS) for the contact dependent translocation of substrate proteins into eukaryotic host cells or are responsible for the survival of salmonellae in macrophages.10 Other virulence factors can be encoded on plasmids. Those factors are associated with enhanced virulence such as the spv cluster that is essential for infection in laboratory rodents. However, the role of virulence plasmids in gastroenteritis and invasive disease in humans is still unclear. Some reports suggest that the spv genes promote dissemination of serotype Typhimurium from the gut and plasmid carrying strains show an increase of the growth rate of the bacteria in liver and spleen.11 Toxins which are released in the environment are not known for Salmonella.

32.1.3 Epidemiology In 2006, a total of 160,649 confirmed cases of human salmonellosis (TESSy) were reported in the EU. The EU incidence was 34.6 cases per 100,000 inhabitants, ranging in the countries from none to 235.9 cases per 100,000. In the United States approximately 40,000 cases of salmonellosis are reported annually.12 It is estimated that the real number of infections exceeds the tenfold. Together with Campylobacter sp. Salmonella is the most widespread foodborne pathogen in Europe. S. enteritidis and S. typhimurium are the most epide­ miological important serovars, because they are responsible for more than 80% of all human infections worldwide.13 In Europe, S. Enteritidis is implicated in over 60% of cases of human salmonellosis.14 Other serovars causing human infections (approx. 1–3%) are S. virchow, S. infantis, S. hadar, S. heidelberg, and S. paratyphi B. However, the prevalence of certain serovars can vary between the countries. The transmission of Salmonella is cyclic between humans, animals, food, and environmental sources. Usually, nontyphoidal salmonellae spread along the food chain. In farm livestock animal feed and high levels of fecal shedding of infected animals has been recognized as important entry site in the food chain. Furthermore, another source of contamination is the slaughtering of the animals. In underdeveloped countries fecal contamination of water is a significant source for S. typhi and S. paratyphi A human infections. The reservoir for S. Enteritidis is mainly poultry often adopting asymptomatic infection in the animals.15 The pandemic of S. enteritidis might have started in the mid 1980s16 and involves interactions of the pathogen with the environment, especially hen house conditions, the birds, the eggs as well as the human host.17 PT4 was in the 1990s formerly the predominant phage type within S. enteritidis.18 However, between 1998 and 2003 a dramatic shift in the proportion of

449

Salmonella

(iv) confirmation of presumptive Salmonella isolates. The whole procedure takes 3–7 days and is very labor-intensive. These four steps are included in standard methods recommended by the US Food and Drug Administration’s (FDA’s) Bacteriological Analytical Manual (BAM), Association of Official Analytical Chemists International (AOACI) as well as the International Organization for Standardization (ISO).27–29 The internationally accepted method for the detection of Salmonella in food and animal feed is described in EN/ISO 6579:2002.28 Although the steps included in the methods for detecting Salmonella spp. approved by different organizations are basically the same, the media used in the different steps differ. The media used is also depending on the type of sample being examined. The outline of the ISO 6579:2002 method is shown in Figure 32.1. Pre-enrichment in a nonselective medium is necessary for efficient recovery of sublethally injured Salmonella in heated, dried, irradiated or otherwise processed samples, such as food and feed.30 In these samples Salmonella is still viable and able to cause disease under the right conditions, but the organisms are easily killed if they are grown under selective pressures such as high temperature or in the presence of chemical additives.31 On the other hand, it has been suggested that the use of nonselective enrichment could mask the presence of Salmonella by allowing the growth of a competing microflora. Buffered peptone water (BPW) and lactose broth are two of the most widely used media for pre-enrichment. The most commonly recommended sample/ pre-enrichment broth ratio is 1:9, i.e., 25 g food to 225 ml broth. The incubation temperature is usually 35–37°C and the incubation time is 18–24 h. Selective enrichment has the purpose of increasing the levels of Salmonella to enable detection on the selective agar plates. This is accomplished by inhibiting the growth of other microflora by the addition of different selective agents in the media. Three of the most commonly used media are Rappaport–Vassiliades soy (RVS) broth, selenite cysteine (SC) broth and tetrathionate (TT) broth. RVS contains malachite

phage types affecting humans in Western Europe was recognized.19 PT4 is replaced by non-PT4 (mainly PT8, PT14b, and PT21) phage types. Other serovars such as S. Paratyphi B have been isolated more frequently within the last years from poultry20,21 and this serovar spread to humans recently causing a multinational outbreak.22 Due to its multiresistance it is regarded as a serious new emerging serovar.14 S. Typhimurium is beside S. Derby frequently isolated from pig and pig products. Most of these strains belong to a special phage type DT104 which emerge since the late 1990s in pigs worldwide.6 DT104 is associated with enhanced virulence and multidrug resistance. This type harbors usually a pentaresistance to ampicillin, chloramphenicol, streptomycin, sulfonamides, and tetracycline (ACSSuT) which is chromosomally encoded by the Salmonella Genomic Island 1 (SGI 1). The size of SGI 1 is 43 kb and originally found in a Canadian S. Typhimurium phage type DT104 isolate but was recently also detected in other, epidemic Salmonella serotypes (e.g., Agona, Albany, Newport, Paratyphi dT + ).23 SGI 1 apparently spread horizontally and is a public health risk in regard to the future treatment of Salmonella infections. Human outbreaks caused by DT104 occurred periodically. Recently, a new multiresistant monophasic serovar (S. 4,12:i:-) is emerging in pig. This serovar has been found initially in Spain24 and spread since then throughout Europe.25

32.1.4 Conventional Diagnostics Salmonella can be isolated by direct plating on selective or nonselective plates from samples such as blood or feces containing high numbers of target bacteria.26 However, food, feed and environmental samples usually contain low numbers of potentially stressed organisms and therefore need a more extensive detection procedure. The conventional routine analysis methods for isolating Salmonella from food, animal feed and environmental samples rely on four basic steps: (i) pre-enrichment in nonselective media, (ii) enrichment in selective media, (iii) isolation on selective agar plates, and Day 0

25 g meat + 225 ml BPW Enrichment (BPW)

Day 1

Enrichment (RVS, MKTTn)

41.5ºC, 24 h,

Centrifugation

37ºC, 24 h,

Day 2

Selective plating (XLD, one optional plate)

Day 3

Bio chemical and serological identification ISO 6579:2002

37ºC (ISO 18 ± 2 h, PCR method 12 ± 2 h)

DNA extraction

PCR

37ºC, 24 h

PCR method

Figure 32.1  The outline of the culture based ISO 6579:2002 method28 and the real-time PCR method95 for the detection of Salmonella in meat samples. The methods are described in more detail in the text.

450

green and magnesium chloride and has a slightly reduced pH; SC contains sodium acid selenite and TT brilliant green (BG) dye and bile. Typically a volume of between 0.1 and 1 ml of the pre-enriched sample is transferred to the 10 ml of selective broth and is incubated at 35 or 42°C depending on type of sample and media used. It has been shown that levels after selective enrichment must exceed 104 CFU/ ml for Salmonella to be detectable on the plates.32 Modified semisolid Rappaport–Vassiliades (MSRV) is another selective media that is particularly useful to detect salmonellae in feces and environmental samples.33 It is based on the ability of salmonellae to migrate through the selective medium ahead of competing motile organisms, thus producing opaque halos of growth. In the third step of the culture based method, a loopful of the selective enrichment broths are streaked on to the surface of selective solid media to obtain isolated colonies. The plates are then incubated at 35–37°C for 18–24 h. The use of plating media serves two basic needs: to selectively inhibit the growth of other bacteria by the addition of inhibitory compounds and to differentiate Salmonella from other bacteria, e.g., by the production of H2S or acid from some sugars. Many different plating media are available and it is recommended that at least two different media be employed since each agar uses one or more different selective agent. Commonly employed media include BG, xylose lysine Tergitol-4 (XLT4), bismuth sulphide (BS), Hektoen enteric (HE), and xylose lysine deoxycholate (XLD) agars. Atypical colonies cause problems in the analysis, both the appearance of Salmonella-like false-positive colonies, e.g., Proteus, and false-negative colonies, e.g., H2S-negative31 and lactosefermenting Salmonella isolates.34 The overgrowth of nonSalmonella strains can mask the presence of Salmonella, thereby making the selection of colonies for further analysis difficult. The final step consists of the identification of the selected isolates by biochemical tests and serotyping. Presumptive positive colonies from the selective agar plates are inoculated onto a nonselective plate in order to get well isolated colonies that can be used for further tests. Several biochemical tests are then employed, i.e., Triple sugar iron (TSI) agar, mannitol, urea, ornithin decarboxylase and lysine decarboxylase.35 Suspect colonies are thereafter verified serologically as Salmonella by determining the antigenic composition. The antigens are classified as somatic (O) and flagellar (H) and are determined by an agglutination test using somatic and flagellar polyvalent antisera. Complete serological typing is performed at the National Reference Laboratories for Salmonella. The use of modified conventional methods, for instance new selective broths or plates, has the advantage of increased selectivity.36 On the other hand, many of these media are expensive and depend on characteristics that are not expressed by all Salmonella strains, such as mobility. For confirmation miniaturized tests, i.e., API 20 E (bioMérieux, Hazelwood, MO) and BBL™ Enterotube™ II (BD Diagnostics, Sparks, MD), have been developed and they provide an efficient

Molecular Detection of Foodborne Pathogens

and labor-saving strategy for identification of suspected Salmonella isolates.37 However, modified conventional methods are still time-consuming techniques whose main advantage is that they provide an isolate that can be used for further characterization. In contrast, nonisolating techniques do not provide an isolate, but since they are faster and more convenient some methods are in use today for the detection of foodborne pathogens. Methods based on conductance or impedance rely on the changes arising from microbial growth in a medium.38 The selectivity of the method depends on the characteristics of the medium and problems associated with false-negative and false-positive results have been reported. The main advantage is the ability to automate the process, and one example is the Malthus 2000 system (Malthus Instruments Ltd., West Sussex, England) that has been used for the detection of Salmonella in food and feed samples.39 In conclusion, the traditional detection methods for Salmonella are slow and labor-intensive. Furthermore, these methods suffer from poor specificity due, for instance, to difficulties in recovering sublethally injured cells or problems in the identification of atypical colonies. These obstacles can be overcome by using molecular methods either in combination with culturing or by direct analysis of the sample. There is a general opinion that the culture-based standard methods are the “golden standard” which should be used in microbiological analysis of foodborne pathogens. To gain acceptance among authorities and end users, it is important that the alternative method be proven to be as good as, or better than, traditional methods.

32.1.5 Molecular Diagnostics Because of the seriousness of Salmonella as a human pathogen much effort has been devoted to the development and improvement of detection methods in various matrices including clinical, environmental, food, and feed samples. These methods should be rapid, sensitive, specific, and user-friendly. The traditional culture based methods do not in fully meet these demands and therefore several methods based on the detection of intracellular macromolecules such as proteins, RNA, and DNA have been developed. To gain acceptance, the methods must give comparable results to those obtained with the traditional culture-based methods. Several molecular methods have been proposed, but since the numbers of Salmonella bacteria in most foods and feeds are small, most methods still rely on conventional culturing methods for enrichment.6 When choosing a method obvious factors such as reliable detection of low amounts of Salmonella, low detection limit, cost, rapidity, ease of handling, possibility of automation, and international acceptability must be taken into consideration.29 Routine analysis of pathogens in food and feed samples places additional demands on the methods used. Apart from the general requirements, such as speed, high sensitivity and specificity, factors such as low cost, the ability to detect low numbers of target microorganisms (the official demands is 1 CFU/25 g feed or food), official approval and lack of false-negative as

451

Salmonella

well as false-positive results are of importance. There is also a growing demand for methods that can be performed at-line or on-line. Many methods that have been developed for clinical applications where, for instance, the requirements on cost and detection level are not as high are, for these reasons, not suitable for routine analysis of Salmonella in food or feed samples. A summary of some of the more commonly used molecular methods for the detection of Salmonella is given in Table  32.1. Molecular methods used for the detection of Salmonella may be divided into two main categories: (i) immunological techniques including enzyme-linked immunosorbent assays (ELISA) and immunomagnetic separation (IMS) and (ii) nucleic acid-based assays including PCR and microarray.29 These two categories will be explained in more detail in the sections below and some examples of methods will be described. 32.1.5.1  Immunological Methods Immunological methods such as ELISA, enzyme-linked fluorescent assays (ELFA) or immunoagglutination are based on the detection of Salmonella specific antigens and have a rather high detection limit (Table 32.1). For this reason an enrichment step is usually applied. Since the specificity depends on the antibodies both false-positive and false-negative results have been reported. However, because of the ease of automation, a number of commercial kits and machines are available, e.g., the VIDAS SLM™ (bioMérieux) and EiaFoss Salmonella™ (Foss Electric Co., Hillerød, Denmark) systems (Table 32.1).41,52 The VIDAS SLM™ Salmonella assay is an automated enzyme immunoassay method which detects Salmonella antigens using the ELFA method on the VIDAS or miniVIDAS analyzers and has been employed for the detection of Salmonella in foods and in agricultural products.53 Before analysis using the VIDAS system, samples are pre-enriched in BPW and enriched further in M broth for a total time of

22–28 h. Presumptive positive results with the VIDAS system need to be culturally confirmed by the isolation and identification of viable salmonellae from the enrichment broths. The Eia–Foss system is an automated fluorescent enzyme immunoassay method for the detection of motile Salmonella in foods, animal feed, and environmental samples.36,54 The system is based on specific binding of salmonellae to monoclonal antibodies for flagellar (H) Salmonella antigens. The surface of paramagnetic microspheres is pre-coated with Salmonella-specific antibodies for the capture of target antigens in the test samples. A portion of boiled M broth post enrichment culture is placed in the sample vial and is then automatically analyzed by the instrument. Salmonella antigens present in a test sample bind to antibodies attached to the beads and unbound sample material is removed by washing. Signal antibodies conjugated with B-galactosidase and a substrate is added after washing steps. The substrate is converted into a fluorescent product by the enzyme conjugate and the intensity of emitted fluorescence is measured by an optical scanner. Presumptive positive results with the EiaFoss Salmonella system need to be culturally confirmed by the isolation and identification of viable salmonellae from the enrichment broths. Two examples of commercially available ELISA kits are the TECRA Salmonella Unique™ test (TECRA, Frenchs Forest, Australia) and the Bioline Selecta (Bioline Ltd, London, UK) which both are performed in the sandwich configuration.55 Salmonella specific antibodies are adsorbed onto the surface of wells where the enriched sample is added. If Salmonella antigens are present in the sample, they are captured by the antibodies. All other material in the sample is washed away. Enzyme labelled antibodies specific for Salmonella are added and the presence of Salmonella is indicated when the bound conjugate converts the substrate and a color change is detected. Presumptive positive Salmonella should be confirmed by streaking from the enrichment broth onto selective agar plates.

Table 32.1 Examples and Characteristics of Different Molecular Methods That Have Been Used for the Detection of Salmonella Category Immunological

Nucleic-acid-based

a  b 

Method

Detection Level (CFU/ml)a

Ease of Handlingb ±

Examples of Commercially Available Methods or Kits

Reference

ELISA/ELFA

103–105

IMS

102–103

±

VIDAS SLM™ EiaFoss™ TECRA Salmonella Unique™ Dynabeads™

Dip-stick

106

 + 

Reveal™

43

Antibody microarray

107

-

DNA hybridization

103–105

±

GeneQuence™

39

DNA microarray

104–107

-

PCR

102–103

 + 

The detection level is the number of Salmonella cells/ml that are needed to yield detection. Ease of handling: +, easy, ±, intermediate, -, difficult.

40,41

36,42 44 45,46

BAX™ TaqMan™ Salmonella Detection kit IQ-Check™ kit

47–51

452

Dip-stick assays are convenient and rapid and one commercially available example is the Reveal system (Neogen, Lansing, MI).56 Reveal contains Salmonella specific antibodies bound to colloidal gold and, separately, to a solid support matrix. Any Salmonella antigen present will bind to the gold conjugated antibodies forming an ­antigen–antibody–chromogen complex. This complex flows across a lateral flow membrane and is subsequently bound by antibody immobilized on the membrane. This causes the gold conjugate to precipitate, forming a visible line and indicating a positive reaction. Proper test completion and flow is indicated by a control line which forms further up in the test window and verifies a valid test run. The time of analysis is only 10 min, but samples need to be enriched in selective media prior to analysis. The more recent advances in immunological techniques include the use of protein microarray, where spots or parallel printed stripes containing antibodies against Salmonella are printed on glass or microtiter plates.44 Gehring et al. reported a sandwich assay for Salmonella with fluorescein- or Cy3labeled reporter antibodies where the capture antibodies were printed within individual wells of streptavidin-coated polystyrene 96-multiwell microtiter plates.44 The detection level was reported to be 107 CFU/ml of enriched sample when tested on a culture-enriched ground beef filtrate and the assay time was 2.5 h. Furthermore, a Salmonella O-antigen microarray was developed by Blixt et al., where oligosaccharide antigens was covalently coupled to a glass or silica chip.57 Antigens specific to serogroups A, B, and D were included in the assay and could be detected in human sera from patients diagnosed with salmonellosis and the authors conclude that this might be a future more rapid, precise, and low-cost alternative for screening of human and livestock for Salmonella infections. 32.1.5.2 Nucleic Acid-Based Methods Nucleic acid-based methods detect DNA or RNA specific for the target organism. Among the nucleic-acid-based methods, PCR is the most widely used. PCR, which is a very powerful tool in molecular biology, mimics nature’s own replication procedure.58 This technique enables the identification and quantification of selected microorganisms and there are several different PCR systems available. The conventional method is based on thermal cycling in a heat block followed by detection by, for instance, electrophoresis using an ethidium-bromide-stained gel59 or immunological techniques such as ELISA.60 To decrease the risk of cross contamination, shorten the analysis time and increase the possibility of automation it is desirable for amplification and detection to be combined in one step, i.e., real-time PCR. Two different detection principles are usually applied in real-time PCR:61 (i) fluorescent dyes that bind specifically to double-stranded DNA (ds-DNA), and (ii) sequence-specific fluorescentlabeled probes. Among the advantages of using ds-DNAbinding dyes are the simplicity in developing such an assay and the lower cost compared with a probe-based assay. On the other hand, due to the detection of unspecific primerdimers in the later cycles, the sensitivity is lower. Because

Molecular Detection of Foodborne Pathogens

of the lower specificity, it may be difficult to show that the right product has been detected and therefore more difficult to introduce such a system for diagnostic use. However, this problem can be overcome to some extent by the use of melting curve analysis. Several probe-based systems are available and among the most commonly used are: hydrolysis probes or TaqMan probes,62 hybridization probes,63 and molecular beacons.64 The advantages of using sequence-specific probes include increased specificity and no need for post-PCR confirmation of the product identity. A large number of PCR based detection methods for Salmonella have been described in literature for clinical as well as for food and feed samples. The majority of these assays amplify a part of the invA gene, encoding a protein involved in the invasion of epithelial cells.51 However, it has been shown that invA is lacking in some strains.51,65 Recently, a PCR assay was designed for amplification of a part of the ttrRSBCA locus encoding proteins used for TT respiration.50 This assay have been validated in collaborative trials and found to perform well.66 Furthermore, a number of validated PCR based systems and kits are commercially available for detection of Salmonella in various matrices (see Table 32.1). One example of a widely used commercially available system employing DNA based detection is the BAX® system (DuPont Qualicon, Wilmington, DE).47 After a 22–26 h pre-enrichment, followed by a 3 h regrowth step or overnight secondary enrichment in TT/RVS in some foods, DNA is extracted and analysed using PCR in an automated procedure. Two examples of commonly used PCR based kits for detection of salmonellae are the TaqMan™ Salmonella kit (Applied Biosystems, Foster City, CA) and the iQ-Check™ Salmonella kit (Bio-Rad Laboratories, Hercules, CA).52,67 In both kits DNA is extracted from a sample pre-enriched in BPW for 16 h. DNA is then analyzed using PCR. DNA can also be detected directly without the use of PCR amplification by employing hybridization techniques. GeneQuence™ (Neogen, Lansing, MI) is a commercially available detection assay employing DNA hybridization technology in a microwell format to detect Salmonella. A  portion of the lysed enrichment culture is hybridized to an oligonucleotide capture probe specific to rRNA sequences and labeled at the 3′ end with polydeoxyadenylic acid (poly dA); and an oligonucleotide detector probe also specific to rRNA sequences of the target organism and labeled at the 5′ end with the enzyme horseradish peroxidase (HRP). Both probes will hybridize to their complementary sequences on the target molecule. The resulting complex is captured onto the solid phase coated with poly deoxythymidylic acid (poly dT), which is complementary to the poly dA portion of the capture probe. Unbound probe is then washed away, and a substrate of HRP is added. A color change is detected if Salmonella is present. An expanding field within the nucleic acid based methods is the use of DNA microarrays for detection and characterization of pathogens such as Salmonella. Microarrays allow the detection of specifically labeled DNAs from many

Salmonella

different target genes simultaneously by binding to oligonucleotides or PCR probes immobilized to small glass slides. Arrays detecting the full genome or specific genes involved in for instance antibiotic resistance and virulence or serotype specific genes have been described.46,68–70 Furthermore, arrays have been developed to detect salmonellae directly in the sample in the presence of other microorganisms (Table 32.1). The detection sensitivity in this case has been reported to be 0.1% (corresponding to 104 genome copies) of the total DNA for the combination of PCR followed by microarray hybridization.71,72

32.2  Methods 32.2.1 Sample Collection and Preparation 32.2.1.1 Sample Collection In many samples and in particular in feed and food samples, the levels of Salmonella are low and unevenly distributed, which means that appropriate sampling and sample preparation steps are crucial to obtain the correct analysis result. Furthermore, the limit of detection in the traditional culturebased method is 1 CFU per 25 g food or feed.28 Although molecular methods such as PCR is sensitive enough to detect one single copy of DNA in the reaction tube, a sample preparation step is needed to reduce the size of the heterogeneous bulk sample to a small homogeneous PCR sample, and to concentrate the small numbers of cells into the size of a PCR sample, i.e., 1–10 μl, or multiply them to a concentration that lies within the practical operating range of the molecular technique applied, i.e., 104 –108 CFU/ml.49 In addition, substances that interfere with the consequent detection method, i.e., PCR, should be removed or neutralized. Four different approaches can be used to withdrawn a representative sample, namely direct, maceration, rinsing, and swabbing.73 The choice of sampling technique depends on several factors such as the sample matrix, level of automation/instrumentation requested as well as the number of samples needed to statistically reflect the original microbial status of the sample. 32.2.1.2 Sample Preparation Since most complex biological samples contain substances that interfere with nucleic acid based or immunological detection methods,74 numerous sample preparation protocols have been developed and optimized. The aim of these procedures is to convert a complex biological sample with its target microorganisms into a sample that is compatible with molecular detection techniques by combining optimal sampling, sample preparation and/or detection conditions.75,76 The reason for the many detection protocols and sample preparation methods employed is that the most suitable strategy depends on the nature of the sample and the purpose of the analysis. Since the numbers of many pathogenic bacteria such as Salmonella are low in many kinds of samples, most of these methods rely on the use of culture enrichment.77 This step is often followed by either DNA extraction or a physical

453

sample preparation method. In general, sample preparation may be divided into four major categories:78 (i) biochemical, (ii) culture enrichment, (iii) immunological, and (iv) physical methods. When choosing a method, the nature of the sample and the aim of the analysis should be considered. Biochemical methods, including direct extraction of nucleic acids, have been used extensively for the detection of Salmonella and other pathogens. The advantage of these methods is that a homogeneous sample of high quality is obtained for PCR amplification and that most PCR inhibitors are removed. The main disadvantage is the interference of the nontarget DNA that is co-purified with the target nucleic acids.79 Culture enrichment prior to detection using molecular techniques includes multiplication of the target microorganism in a selective or nonselective liquid medium. The use of culture enrichment prior to molecular analysis serves many purposes, including: the dilution of inhibitory substances present in the sample matrix, multiplication of the target organism to provide detectable concentrations, dilution of dead cells, and last, but not least, the possibility of isolating the target organism for complementary tests.80 The specificity will depend on the characteristics of the medium. If the medium is nonselective, such as BPW, growth of background flora may occur and disturb the detection since DNA or cells other than the target organism have been shown to affect both the sensitivity and specificity.79 Most enrichment media contain substances that will interfere with PCR amplification or immunological methods and therefore the choice of medium is of importance. Immunological methods are based on the detection of specific antibodies and several basic techniques are available. Magnetic beads coated with antibodies are used extensively for the detection of foodborne pathogens such as Salmonella,81 in a technique referred to as IMS. There are also several automated systems available for detection of these bacteria and other pathogens.41,81 Immunological methods are relatively easy to automate and are therefore often used for routine analysis purposes. However, the specificity of the assay depends in part on the antibodies used and the binding of the antibodies has been shown to be affected in biological samples such as feces.82,83 Physical sample preparation methods, such as aqueous two-phase systems,84 buoyant density centrifugation,85,86 centrifugation,87 dilution,49,88 filtration,89 and flotation,86,90 depend on the physical properties of the target cell, e.g., cell density and size. These methods have been extensively applied in the detection of foodborne pathogens. The main advantage with physical methods is that they do not influence the specificity, as is the case with immunological and culture-based methods. One promising physical method is flotation which is based on a discontinuous density gradient separation technique based on the buoyant density of bacteria.91 In flotation, the sample, e.g., a food homogenate, is loaded at the bottom of a test tube together with a density medium, e.g., BactXtractor™ (QRAB, Bålsta, Sweden). Two or more layers with decreasing densities are applied in layers

454

Molecular Detection of Foodborne Pathogens

on top of the sample. During centrifugation at a moderate g-force cells or particles will float to a level with the same density as their own.92 The recovered target bacterium, separated from inhibitors and matrix particles, can now be used for further analyses. This technique has been used, e.g., as a pre-PCR treatment method to separate pathogenic bacteria such as Salmonella from food86,90 and to separate living and dead Campylobacter cells.93 Furthermore, it has been found that flotation can be applied as a sample preparation method prior to PCR, to avoid the detection of DNA originating from dead cells.94 Since the flotation method is nondestructive, cells can be recovered for further identification and characterization, which is advantageous when, e.g., performing epidemiological investigations. Another advantage of using flotation lies in the possibility of scaling up and automation of the process. However, the method needs to be standardized and each target bacterium/species requires a specific protocol. In conclusion, using flotation as a sample preparation method in combination with PCR provides the possibility of future, real-time at-line detection of living Salmonella in food and animal feed samples.

32.2.2  Detection Procedures Principle: The method described here is based on a shortened pre-enrichment in BPW followed by automated DNA extraction and detection by a 5’ nuclease (TaqMan) real-

time PCR.50,95,96 It has been validated and approved by the Nordic Organization for Validation of Alternative Methods (NordVal) for qualitative analysis of salmonellae in raw meat samples.97,98 The assay uses specifically designed primers and a probe targeting within the ttrRSBCA locus,50 which is located near the Salmonella pathogenicity island 2 at centisome 30. Both the inclusivity and exclusivity of the primers have been determined to 100%.50 The total analysis time is 14 h and the detection limit is 1–10 CFU per 25 g raw meat. The method has also been evaluated for other matrices, i.e., carcass swabs and animal feces and approved by NordVal.97 Reagents: The following reagents are needed for automated DNA extraction: (i) Magnesil KF, Genomic System (Promega Corporation, Madison, WI; store at room temperature); (ii) ethanol (95–100%, room temperature); and (iii) isopropanol (room temperature). Reagents for real-time PCR are listed in Table 32.2. Apparatus and equipment: Mx3005p Real-time PCR thermal cycler (Stratagene, La Jolla, CA), or equivalent equipment; plastic ware for real-time PCR; KingFisher (Thermo Fisher Scientific, Waltham, MA) or equivalent equipment; plastic ware for DNA extraction; centrifuge for 5 ml samples (3,000 × g); micropipettes: 20 µl, 100 µl, 1 ml, and 5 ml; filter tips for micropipettes, PCR grade; Eppendorf tubes: 0.5 and 1.5 ml, PCR grade; incubator, 37.0±1.0°C; and bottles, 100–500 ml.

Table 32.2 Reagents Used in the PCR Mastermix of the TaqMan Real-Time PCR Method for Detection of Salmonella in Meat Described in the Text Reagent

Volume/Sample Concentration of Stock (µl) Solution

PCR H2O PCR buffer

0.25 2.5

dUTP mix MgCl2 Glycerol Primer 1

1.0 4.0 2.0 1.0

Primer 2

1.0

BSA

Sequence or other Relevant Information

– 10 ×  12.5 mM 25 mM 87%

10 × buffer: 100 mM Tris-HCl pH 8.9 (25°C), 1 M KCl, 15 mM MgCl2, 500 µg/ml BSA, 0.5 % Tween 20 (v/v) 2.5 mM each of dATP, dCTP, dGTP, and 5.0 mM of dUTP

10 pmol/µl

5´ CTC ACC AGG AGA TTA CAA CAT GG 3´ 5´ AGC TCA GAC CAA AAG TGA CCA TC 3´

1.25

10 pmol/µl 20 mg/ml

Tth DNA polymerase DMSO Salmonella LNA probe

0.3

5 U/µl

0.5 0.6

10 pmol/µl

5′-(6FAM) CG + ACGGCG + AG + ACCG (BHQ1)-3′ ( + indicates LNA substitution)

IAC probe

0.6 1.0

10 pmol/µl a

5′-(JOE) CAC ACG GCG ACG CGA ACG CTT T (BHQ1)-3′

IAC target Template DNA Total volume/sample

9.0 25.0

– –

a

Dissolved in 50 mM Tris-HCl, 100 mM NaCl, 0.25 mM EDTA, 1 mM 2-mercaptoethanol, 50% glycerol (v/v), pH 7.5

Dimethyl sulfoxide, PCR Reagent, SIGMA D9170-1VL or equivalent

5′-GACTCACCAGGAGATTACAACATGGCTCTTGCTGTGCATCATCGCAGAAC ATCAAAGCGTTCGCGTCGCCGTGTGGGATGGTCACTTTTGGTCTGAGCTAC-3′

The concentration used should result in a Ct value in the HEX channel of 28–32.

Salmonella

455

Procedure:

Reporting of results: A sample is considered positive if a Ct value for FAM below 36 is obtained, and the controls are as expected. Due to competition of amplification between the Salmonella target and the internal amplification control, absence of signal from the IAC can occur in strong positive samples. A sample is considered negative if no Ct value for FAM is obtained during the 40 cycles (no amplification has occurred) or a Ct > 36 is obtained and a Ct value for JOE/HEX below 40 is obtained. If the IAC is not amplified in negative samples, the result may be false negative and the analysis should be repeated.

(1) Weigh 25 g of raw meat in a stomacher bag and pre-warm the samples to 37°C ± 1.0°C. Add 225 ml 37 ± 1.0°C BPW. Homogenize manually for 5 sec and incubate at 37°C ± 1.0°C for 12 ± 2 h. Remove 5 ml for DNA extraction. Centrifuge the sample at 3,000 × g for 5 min, and discard the supernatant. Keep the sample pellet at –20.0°C ± 2°C until performing DNA extraction. Perform automated DNA extraction using the KingFisher and the MagneSil KF Genomic System or equivalent as previously described.95 Re-suspend the sample pellet in 200 µl of lysis buffer and prepare the plates for the automated DNA extraction (75 µl of paramagnetic beads is used). Place the prepared plates in the automated DNA extraction platform, and run the program. Use 9 µl of the extracted DNA as template in the realtime PCR. (2) Prepare PCR master mix using reagents and volumes shown in Table 32.2. Prepare a batch of PCR master mix large enough to analyze all samples in the analysis to avoid intra analysis variations. Always prepare a small excess of master mix to ensure a sufficient amount for all samples. Dispense 16 µl of the mix per tube in the real-time PCR tubes, and add 9 µl of the extracted DNA. Include controls as described below. (3) The controls are: (i) Incubation control: Salmonella spp. in 250 µl BPW incubated with the samples; (ii) DNA extraction controls: A process blank (lysis buffer), and a positive control to check the DNA extraction (dilute enriched culture in 0.9% NaCl solution until approx 40,000 salmonellae/ml and dispense 50 µl into 200 µl lysis buffer); (iii) PCR controls: An internal amplification control (IAC) to detect false negative responses, a positive DNA control (Salmonella DNA in a concentration of approximately 0.005 ng), a nontemplate control (NTC, only the master mix and PCR grade water) and a negative DNA control (Escherichia coli DNA in a concentration of approximately 5 ng). (4) Place the PCR tubes with the mix and added samples and controls and run the following thermal profile: one cycle of 95°C for 3 min; 40 cycles of 95°C for 30 sec, 65°C for 60 sec, and 72°C for 30 sec. Note: The thermal cycler should be in compliance with the requirements for installation, performance, and maintenance described in ISO/TS 20836.99 Fluorescence measurements are obtained online and should be analyzed with the appropriate software. The threshold should be set as: “Backgroundbased threshold” using Mx Real-time PCR thermal cycler (Stratagene). Data should be collected in the annealing step from the probe fluorophores FAM (emission 520 nm) and JOE (emission 550 nm, use the “HEX” channel) in the realtime MX PCR thermal cycler.

32.3 Conclusions and Future Perspectives A large number of molecular methods, including immunological and nucleic acid based methods, for detection of Salmonella in various matrices have been published. Furthermore, several kits and equipment are commercially available for this purpose. However, there are still some challenges left before molecular methods will replace traditional culture based methods. One challenge facing users of diagnostic PCR on a routine basis for the analysis of foodborne pathogens such as Salmonella, is the low number of standardized, officially approved methods. This has previously limited the use of PCR for other kinds of routine analysis. Validation is an important step in the process of standardizing a method, because it provides evidence that the new method gives equivalent results to the currently used reference methods, as well as providing confirmation of the reproducibility and specificity when used by other laboratories.100 These data are needed to gain acceptance among authorities and end users for the new method and to speed up the implementation of new rapid PCR-based detection systems at diagnostic analysis laboratories. A lot of the molecular methods available in literature have not been validated in comparative and collaborative studies. An important limitation with current molecular techniques lies in the detection time. Due to the low numbers of salmonellae present in particularly food, feed, and environmental samples an enrichment step is needed to yield numbers high enough (approximately 104 CFU/ml) to enable detection using molecular methods. This slows down the analysis; the time for molecular analysis is usually between a few minutes up to four hours, compared to enrichment times of at least 8 h. Thus, there is a need to develop efficient methods that enable detection of low numbers of cells directly in the sample. Besides the challenge in food laboratories, clinical laboratories are facing a similar challenge to food laboratories; increasing number of human stool samples are submitted at late excretion phase and contain low number of targets. The situation is worse in veterinary laboratories, where fecal samples, especially from intermediate carrier cows, can contain very low number of S. Dublin. Not to mention feed samples containing substantial plant ingredients, all strongly

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inhibitory to PCR, or other detection methods. Thus, the main future focus should be on more efficient, and at the same time simpler, pre-analytical and quantitative sample preparation methods. Quantitative methods are needed because we need a better picture of how many target genomes were present in the initial sample, how many were lost due to sample preparation, and what is being detected in the final assay. The way we evaluate the current methods is just by a side-by-side comparison with culture methods, but this does not provide us with a clear understanding of methods. It is tumbling in dark. In addition, quantitative risk assessments require quantitative detection in order to produce meaningful numbers for decision-support modeling studies. Another important future focus is faster characterization and (sero)-genotyping of isolates; new microfluidic technologies make it possible to combine detection and characterization in one assay. The choice of target sequences requires international validation of consensus probe sequences against epidemiologically relevant and well-balanced strains. Studies should look into the cons and pros of mirroring current Kaufmann–White serotyping scheme, especially its use for risk assessments. A one-assay combination of detection and subtyping can provide clinical and food laboratories with a better incitement to replace culture with molecular methods.



Acknowledgments



This work was financially supported by the European Union project BIOTRACER (FOOD-2006-CT-036272), the Danish Directorate for Food, Fisheries, and Agri-Business (DFFE) grant 3414-04-01032 and the Swedish Research Council for Environment, Agricultural Sciences and Spatial Planning (FORMAS).

References



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457 48. Hoorfar, J., Ahrens, P. and Rådström, P. Automated 5' nuclease PCR assay for identification of Salmonella enterica. J. Clin. Microbiol., 38, 3429, 2000. 49. Löfström, C. et al. Rapid and specific detection of Salmonella spp. in animal feed samples by PCR after culture enrichment. Appl. Environ. Microbiol., 70, 69, 2004. 50. Malorny, B. et al. Diagnostic real-time PCR for detection of Salmonella in food. Appl. Environ. Microbiol., 70, 7046, 2004. 51. Rahn, K. et al. Amplification of an invA gene sequence of Salmonella typhimurium by polymerase chain reaction as a specific method of detection of Salmonella. Mol. Cell. Probes, 6, 271, 1992. 52. Uyttendaele, M., Vanwildemeersch, K. and Debevere, J. Evaluation of real-time PCR vs automated ELISA and a conventional culture method using a semi-solid medium for detection of Salmonella. Lett. Appl. Microbiol., 37, 386, 2003. 53. Curiale, M.S., Gangar, V. and Gravens, C. VIDAS enzymelinked fluorescent immunoassay for detection of Salmonella in foods: Collaborative study. J. AOAC Int., 80, 491, 1997. 54. Krusell, L. and Skovgaard, N. Evaluation of a new semiautomated screening method for the detection of Salmonella in foods within 24 h. Int. J. Food Microbiol., 20, 123, 1993. 55. Hughes, D. et al. Comparison of the Tecra Salmonella visual immunoassay and standard cultural methods for the detection of salmonellae in foods. Food Technol. Australia, 39, 446, 1987. 56. Bird, C.B., Miller, R.L. and Miller, B.M. Reveal for Salmonella test system. J. AOAC Int., 82, 625, 1999. 57. Blixt, O. et al. Pathogen specific carbohydrate antigen microarrays: a chip for detection of Salmonella O-antigen specific antibodies. Glycoconj. J., 25, 27, 2008. 58. Mullis, K.B. and Faloona, F.A. Specific synthesis of DNA in vitro via a polymerase-catalyzed chain reaction. Methods Enzymol., 155, 335, 1987. 59. Sambrook, J., Maniatis, T. and Fritsch, E.F. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY, 1989. 60. Perelle, S. et al. Comparison of PCR-ELISA and LightCycler real-time PCR assays for detecting Salmonella spp. in milk and meat samples. Mol. Cell. Probes, 18, 409, 2004. 61. Bustin, S.A. Quantification of mRNA using real-time reverse transcription PCR (RT-PCR): Trends and problems. J. Mol. Endocrinol., 29, 23, 2002. 62. Holland, P.M. et al. Detection of specific polymerase chain reaction product by utilizing the 5' to 3' exonuclease activity of Thermus aquaticus DNA polymerase. Proc. Natl. Acad. Sci. USA, 88, 7276, 1991. 63. Woo, T.H. et al. Identification of pathogenic Leptospira genospecies by continuous monitoring of fluorogenic hybridization probes during rapid-cycle PCR. J. Clin. Microbiol., 35, 3140, 1997. 64. Tyagi, S. and Kramer, F.R. Molecular beacons: Probes that fluoresce upon hybridization. Nat. Biotechnol., 14, 303, 1996. 65. Galán, J.E. and Curtiss, R. III, Distribution of the invA, -B, -C, and -D genes of Salmonella typhimurium among other Salmonella serovars: invA mutants of Salmonella typhi are deficient for entry into mammalian cells. Infect. Immun., 59, 2901, 1991. 66. Malorny, B. et al. Multicenter validation study of two blockcycler- and one capillary-based real-time PCR methods for the detection of Salmonella in milk powder. Int. J. Food Microbiol., 117, 211, 2007.

458 67. Chen, S. et al. The evaluation of a fluorogenic polymerase chain reaction assay for the detection of Salmonella species in food commodities. Int. J. Food Microbiol., 35, 239, 1997. 68. Wattiau, P. et al. Evaluation of the Premi(R)Test Salmonella, a commercial low-density DNA microarray system intended for routine identification and typing of Salmonella enterica. Int. J. Food Microbiol., 123, 293, 2008. 69. van Hoek, A.H. et al. Detection of antibiotic resistance genes in different Salmonella serovars by oligonucleotide microarray analysis, J. Microbiol. Methods, 62, 13, 2005. 70. Porwollik, S. and McClelland, M., Determination of the gene content of Salmonella genomes by microarray analysis. Methods Mol. Biol., 394, 89, 2007. 71. Kostic, T. et al. A microbial diagnostic microarray technique for the sensitive detection and identification of pathogenic bacteria in a background of nonpathogens. Anal. Biochem., 360, 244, 2007. 72. Maynard, C. et al. Waterborne pathogen detection by use of oligonucleotide-based microarrays. Appl. Environ. Microbiol., 71, 8548, 2005. 73. Rådström, P. et al. Pre-PCR processing: Strategies to generate PCR-compatible samples. Mol Biotechnol, 26, 133, 2004. 74. Wilson, I.G. Inhibition and facilitation of nucleic acid amplification. Appl. Environ. Microbiol., 63, 3741, 1997. 75. Rådström, P. et al. Optimisation strategies for overcoming PCR inhibition. In PCR Primer: A Laboratory Manual, Dieffenbach, C. and Dveksler, G. Eds. Cold Spring Harbor Laboratory Press, NY, 2003. 76. Rådström, P. et al. Pre-PCR processing of samples. In PCR Detection of Microbial Pathogens: Methods and Protocols, Sachse, K. and Frey, J., Eds. Humana Press, Totowa, NJ, 2003. 77. Stevens, K.A. and Jaykus, L.A. Bacterial separation and concentration from complex sample matrices: a review. Crit. Rev. Microbiol., 30, 7, 2004. 78. Lantz, P.G. et al. Biotechnical use of polymerase chain reaction for microbiological analysis of biological samples. Biotechnol. Annu. Rev., 5, 87, 2000. 79. Wang, R.F., Cao, W.W. and Johnson, M.G. 16S rRNA-based probes and polymerase chain reaction method to detect Listeria monocytogenes cells added to foods. Appl. Environ. Microbiol., 58, 2827, 1992. 80. Sharma, V.K. and Carlson, S.A. Simultaneous detection of Salmonella strains and Escherichia coli O157:H7 with fluorogenic PCR and single-enrichment-broth culture. Appl. Environ. Microbiol., 66, 5472, 2000. 81. Cudjoe, K.S., Krona, R. and Olsen, E. IMS: A new selective enrichment technique for detection of Salmonella in foods. Int. J. Food Microbiol., 23, 159, 1994. 82. Eriksson, E. and Aspán, A. Comparison of culture, ELISA and PCR techniques for Salmonella detection in faecal samples for cattle, pig, and poultry. BMC. Vet. Res., 3, 21, 2007. 83. Nilsson, H.-O. et al. Immunomagnetic bead enrichment and PCR for detection of Helicobacter pylori in human stools. J. Microbiol. Methods, 27, 73, 1996.

Molecular Detection of Foodborne Pathogens 84. Lantz, P.G. et al. Enhanced sensitivity in PCR detection of Listeria monocytogenes in soft cheese through use of an aqueous two-phase system as a sample preparation method. Appl. Environ. Microbiol., 60, 3416, 1994. 85. Shieh, Y.C., Calci, K.R. and Baric, R.S. A method to detect low levels of enteric viruses in contaminated oysters. Appl. Environ. Microbiol., 65, 4709, 1999. 86. Wolffs, P.F. et al. Simultaneous quantification of pathogenic Campylobacter and Salmonella in chicken rinse fluid by a flotation and real-time multiplex PCR procedure. Int. J. Food Microbiol., 117, 50, 2007. 87. Gerritsen, M.J. et al. Sample preparation method for polymerase chain reaction-based semiquantitative detection of Leptospira interrogans serovar hardjo subtype hardjobovis in bovine urine. J. Clin. Microbiol., 29, 2805, 1991. 88. Abu Al-Soud, W. et al. A sample preparation method which facilitates detection of bacteria in blood cultures by the polymerase chain reaction. J. Microbiol. Methods, 32, 217, 1998. 89. Wolffs, P.F. et al. Direct quantitation and detection of salmonellae in biological samples without enrichment, using twostep filtration and real-time PCR. Appl. Environ. Microbiol., 72, 3896, 2006. 90. Wolffs, P. et al. Rapid quantification of Yersinia enterocolitica in pork samples by a novel sample preparation method, flotation, prior to real-time PCR. J. Clin. Microbiol., 42, 1042, 2004. 91. Pertoft, H. et al. The viability of cells grown or centrifuged in a new density gradient medium, Percoll(TM). Exp. Cell. Res., 110, 449, 1977. 92. Pertoft, H. Fractionation of cells and subcellular particles with Percoll. J. Biochem. Biophys. Methods, 44, 1, 2000. 93. Wolffs, P. et al. Quantification of Campylobacter spp. in chicken rinse samples by using flotation prior to real-time PCR. Appl. Environ. Microbiol., 71, 5759, 2005. 94. Wolffs, P., Norling, B. and Rådström, P. Risk assessment of false-positive quantitative real-time PCR results in food, due to detection of DNA originating from dead cells. J. Microbiol. Methods, 60, 315, 2005. 95. Josefsen, M.H. et al. Optimization of a 12-hour TaqMan PCRbased method for detection of Salmonella bacteria in meat. Appl. Environ. Microbiol., 73, 3040, 2007. 96. Reynisson, E. et al. Evaluation of probe chemistries and platforms to improve the detection limit of real-time PCR. J. Microbiol. Methods, 66, 206, 2006. 97. Anonymous. NordVal certificate no 031, NordVal/NMKL, Søborg, Denmark, 2007. 98. Qvist, S. NordVal: A Nordic system for validation of alternative microbiological methods. Food Control, 18, 113, 2007. 99. Anonymous. Microbiology of food and animal feeding stuffs–Polymerase chain reaction (PCR) for the detection of food-borne pathogens–Performance testing for thermal cyclers, ISO/TS 20836:2005. International Organisation for Standardization, Geneva, Switzerland, 2005. 100. Malorny, B. et al. Standardization of diagnostic PCR for the detection of foodborne pathogens. Int. J. Food Microbiol., 83, 39, 2003.

33 Serratia

Zhi-Qing Hu, Wei-Hua Zhao

Showa University School of Medicine

Zhuting Hu

International Christian University

Contents 33.1 Introduction.................................................................................................................................................................... 459 33.1.1 Classification and Pathogenicity....................................................................................................................... 459 33.1.2 S. marcescens as a Common Opportunistic Pathogen..................................................................................... 460 33.1.3 Mechanisms of S. marcescens Resistance to β-Lactams................................................................................. 461 33.1.3.1 β-Lactamases .................................................................................................................................. 461 33.1.3.2 The Outer Membrane Barrier.......................................................................................................... 462 33.1.4 Mechanisms of S. marcescens Resistance to Fluoroquinolones ..................................................................... 462 33.1.4.1 Mutations of gyrA............................................................................................................................ 462 33.1.4.2 Active Efflux Pump.......................................................................................................................... 462 33.1.4.3 Qnr Determinants ........................................................................................................................... 464 33.1.5 Mechanisms of S. marcescens Resistance to Aminoglycosides . .................................................................... 464 33.1.5.1 Aminoglycoside N-acetyltransferases (AAC) . ............................................................................... 464 33.1.5.2 Aminoglycoside O-adenylyltransferases (ANT) ............................................................................ 465 33.1.5.3 Aminoglycoside O-phosphotransferases (APH) . ........................................................................... 465 33.1.5.4 Bifunctional Aminoglycoside Resistance Enzyme.......................................................................... 465 33.1.6 Laboratory Identification of S. marcescens...................................................................................................... 465 33.2 Methods.......................................................................................................................................................................... 465 33.2.1 PCR and Restriction Enzyme Digestion........................................................................................................... 466 33.2.2 Real-Time PCR................................................................................................................................................. 466 33.3 Conclusions and Future Perspectives............................................................................................................................. 466 References.................................................................................................................................................................................. 466

33.1  Introduction 33.1.1 Classification and Pathogenicity Serratia is a genus of bacteria classified as a member of the Enterobacteriaceae family (Table 33.1).1 The bacteria in this family have the following characteristics: (i) they are Gram-negative rods; (ii) they are facultatively anaerobic; (iii) they grow on peptone or meat extract media without other supplements; (iv) they ferment rather than oxidize glucose; (v) they are catalase-positive and oxidase-negative; and (vi) they reduce nitrate to nitrite. S. marcescens is responsible for > 90% of Serratia infections.2–4 The S. liquefaciens complex (S. liquefaciens sensu stricto, S. proteamaculans, and S. grimesii) can also infect humans, while rare species such as S. plymuthica, S. rubidaea, and S. odorifera have also caused human infections.1,2,5 The ubiquitous presence of S. marcescens in the environment, such as in soil, water, air, plants, and animals, is well documented.1 When the temperature and humidity are high, a pinkish film can be seen on bathroom fixtures, sinks or

along the water line of the toilet bowl. This film is most likely from S. marcescens. Historically, the miraculous appearance of ‘blood’ on bread was recorded as early as the sixth century B.C, and often regarded as a religious mystery. In 1823 Bartolomeo Bizio, a bacteriologist at the University of Padua, Italy, revealed that the “blood” was an illusion caused by a living organism. He named this organism S. marcescens.6 The genus Serratia was named in honor of an Italian physicist, Serafino Serrati, who was engaged in inventing a steamboat; and the species marcescens was named from the Latin word “to decay,” following Dr. Bizio’s observation that the red pigment changed color.6 Because S. marcescens is so common and was thought to be nonpathogenic, until the 1960s scientists and teachers, taking advantage of the bright red pigment, frequently used it as a biomarker to trace transmission of bacteria through speech and contact, and to demonstrate the importance of hand washing in schools.6 In the 1950s, the US Army even used it in a secret biological warfare test “Operation SeaSpray” to study how wind might carry biological weapons.7 459

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Table 33.1 Classification of Serratia Domain Phylum Class Order Family Genus Species

Bacteria Proteobacteria Gamma Proteobacteria Enterobacteriales Enterobacteriaceae Serratia    entomophila    ficaria    fonticola    grimesii    liquefaciens*    marcescens    marinorubra ( = rubidaea)    odifera    odorifera    plymuthica    proteamaculans    rubidaea ( = marinorubra)    ureilytica

* Formerly named Enterobacter liquefaciens, this is a heterogeneous species which may be split into three: S. liquefaciens sensu stricto, S. proteamaculans, and S. grimesii.

S. marcescens was released by bursting balloons filled with the bacterium over urban areas of the San Francisco Bay Area in California. Shortly thereafter, an increase in pneumonia and urinary tract infections was observed, although there was no direct evidence that the released strain caused the infections.8,9 In fact, S. marcescens rarely causes primary ­infections in immunocompetent people.1 It is mainly associated with ­hospital-acquired infections in immuno-compromised patients, i.e., it is an opportunistic nosocomial pathogen. S. marcescens is not a common component of the normal bacterial flora of the human gut, and most infections are acquired exogenously.10 Patients can be infected by direct contact, by inhalation of infectious droplets, and by ingestion. S. marcescens can be transferred by contaminated persons, medical devices, intravenous fluids, or other solutions. Just like other nosocomial infections, risk factors include compromised/suppressed immunity, recent surgery, recent corticosteroid therapy, diabetes, cancer, burns, and chronic obstructive pulmonary disease. The very young and the very old are at higher risk.

33.1.2 S. marcescens as a Common Opportunistic Pathogen Among hospitalized patients, nosocomial infections and antibiotic-resistant organisms are two common problems. Nosocomial infections develop in at least 5% of hospitalized patients in the U.S.11 There are more than 2 million nosocomial infections each year, of which 50–60% are caused by antibiotic-resistant organisms.12 Resistance to

antibiotics has become an urgent world-wide problem because it increases the morbidity, mortality and costs associated with nosocomial infections. When inappropriate empirical antibiotic therapy was given to intensive-care units (ICU&) patients infected by resistant bacteria, mortality rates doubled from 30% to 60%.13 As an opportunistic pathogen, S. marcescens is usually associated with urinary and respiratory tract infections. It also causes pneumonia, bacteremia/septicemia, endocarditis, osteomyelitis, conjunctivitis, wound infections, eye infections and meningitis. Sporadic infection is most common, but occasional epidemics and common-source outbreaks occur.2 S. marcescens is isolated with increasing frequency from respiratory specimens, blood, urine, feces and wound exudates of nosocomial patients and is becoming an important nosocomial pathogen, causing outbreaks in nurseries, ICUs and renal dialysis units. It accounts for as much as 1–3% of all nosocomial infections and these are often severe or fatal.2 In 2006, the Japanese Ministry of Health, Labor and Welfare analyzed the data of 170,090 blood specimens from 213 medical facilities. Bacteria were isolated from 15.1% of samples, and 1% were S. marcescens.14 A similar study of 49 hospitals in the United States during 1995–2002, showed that 1.7% of 24,179 nosocomial bloodstream infections were caused by S. marcescens.15 It is important to recognize that the predominant pathogens may vary between geographical regions, between hospitals, and even between units within an individual hospital. In immuno-compromised patients, infections are very difficult to treat effectively, even when the pathogen is sensitive to antibiotics. Nosocomial infections caused by resistant organisms are even more challenging. Unfortunately, this is a particularly serious problem with S. marcescens which has powerful and adaptable resistance. It is often multiply resistant to aminoglycosides, penicillins, cephalosporins and even carbapenems, because of the presence of various resistance mechanisms, and transmissible plasmids (R-factors) carrying one or more resistance genes. Table 33.2 shows the rates of resistance in S. marcescens isolated in Japan during 2006.14 A high proportion of isolates are resistant to ampicillin (89.1%), amoxicillin/clavulanic acid (95.3%), cefaclor (100%), cefazolin (98.5%) and cefpodoxime proxetil (66.3%). The Sanford Guide to Antimicrobial Therapy recommends the third-generation cephalosporins, carbapenems (imipenem/cilastatin, meropenem, and ertapenem) and fluoroquinolones as the first choice antibiotics; aztreonam and gentamicin as the second choice; and ticarcillin/clavulanate and piperacillin/tazobactam as optional agents.16 Over time, treatment becomes more and more difficult because many strains that are initially susceptible to an antibiotic develop resistance during therapy. For example, the effectiveness of the first choice antibiotics, e.g., imipenem and meropenem, is now threatened by the appearance of resistant isolates (4.6% and 1.2%, respectively, Table 33.2). Accordingly, recent research on S. marcescens has focused on the underlying mechanisms of resistance. The major mechanisms by which bacteria exhibit resistance to antibiotics include: drug inactivation or modification by

461

Serratia

Table 33.2 Occurrence of Antimicrobial Resistance Among S. marcescens Isolated from Blood Specimens in 213 Medical Facilities of Japan During 2006 Susceptibility (% of isolates) and MIC (µg/ml)

No. of Isolates Studied

Susceptible

Aztreonam

227

90.7 ≤ 8 

3.5

5.7 ≥ 32 

Ampicillin

247

6.1 ≤ 8 

4.8

89.1 ≥ 32 

Piperacillin

265

76.2 ≤ 16

14.0

9.8 ≥ 128 

Amoxicillin/Clavulanic acid

148

0.6 ≤ 4 

4.1

95.3 ≥ 16 

40

17.5 ≤ 4 

5.0

77.5 ≥ 16 

260

95.4 ≤ 4

0

4.6 ≥ 16 

83

98.8 ≤ 4 

0

1.2 ≥ 16 

Antimicrobial Agent

Ampicillin/Sulbactam Imipenem/Cilastatin Meropenem

Intermediate

Resistant

Cefaclor

152

0 ≤ 8

0

100 ≥ 32 

Cefazolin

265

0 ≤ 8

1.5

98.5 ≥ 32 

Cefpodoxime proxetil

101

32.7 ≤ 2 

1.0

66.3 ≥ 8 

Cefmetazole

221

79.7 ≤ 16

10.8

9.5 ≥ 64 

Cefotaxime

231

77.0 ≤ 8 

11.3

11.7 ≥ 64 

Ceftazidime

261

83.2 ≤ 8 

6.1

10.7 ≥ 32 

Cefepime

84

97.6 ≤ 8

2.4

0 ≥ 32

Amikacin

261

98.1 ≤ 16

0

1.9 ≥ 64

Gentamicin

258

99.6 ≤ 4 

0

0.4 ≥ 16 

Tobramycin

35

82.9 ≤ 4 

5.7

11.4 ≥ 16 

Minocycline

259

78.7 ≤ 4 

15.1

6.2 ≥ 16 

Levofloxacin

262

91.2 ≤ 2 

3.8

5.0 ≥ 8 

74

66.2 ≤ 1 

23.0

10.8 ≥ 4 

199

100 ≤ 38

0

Ciprofloxacin Sulfamethoxazole-trimethoprim

producing a specific enzyme; reduced drug accumulation, either by decreasing drug permeability, or by increasing active efflux of drug; and alteration of target site by gene mutation.

33.1.3 Mechanisms of S. marcescens Resistance to b-Lactams β-Lactams are the most widely used antibiotics in clinics and can be classified as penicillins, cephalosporins, cephamycins, monobactams, and carbapenems, which have a common four-atom structure known as the β-lactam ring. β-Lactams act on transpeptidase and carboxypeptidase (so-called penicillin binding proteins, PBPs) to inhibit the synthesis of the bacterial cell wall. The predominant mechanisms of S. marcescens resistance are the production of β-lactamases, and reduction of outer membrane permeability. 33.1.3.1  β-Lactamases β-Lactamases (EC 3.5.2.6) hydrolyze the β-lactam ring, thus inactivating the drug’s ability to bind with PBPs. More than 500 kinds of β-lactamases have been identified in Gramnegative bacilli (Table 33.3).17 Based on nucleotide and amino acid sequences, they are divided into four classes, A–D (Ambler’s classification).18 Classes A, C, and D belong to serine type enzymes possessing a serine ­moiety at the

0 ≥ 76

active site, whereas class B (metallo-β-lactamases) requires divalent cations, usually zinc, as metal cofactors.18,19 Most β-lactamases are encoded either on chromosome or on plasmid, but overlaps can be observed. Their expression may be inducible, low-level constitutive or high-level constitutive, depending on species and strains.20 S. marcescens naturally produces the class C β-lactamase (AmpC enzyme) encoded chromosomally.21,22 Plasmid-mediated β-lactamases are an increasing threat to treatment, owing to the horizontal transfer of the resistant genes between species.23–25 Two approaches have been employed to conquer β-lactamases: the combination of β-lactam with β-lactamase inhibitor; and development of β-lactams resistant to β-lactamases. However, bacterial strains producing inhibitorresistant TEM enzymes have emerged recently, and this could be related to the frequent use of β-lactamase inhibitors such as clavulanic acid, sulbactam, and tazobactam.26 Carbapenems are highly resistant to most β-lactamases including extendedspectrum β-lactamases, and are effective against resistant bacteria. However, the emergence of carbapenem-hydrolyzing β-lactamases limits their use.27,28 Class A (serine type) and class B (metallo-type) carbapenemases have been found in S. marcescens.29,30 SME-1 is a class A carbapenemase, first identified from two S. marcescens strains isolated in London in 1982.31,32 SME-2 and SME-3, point-mutant derivatives of SME-1, were identified from isolates in the USA.33,34 Although

462

Molecular Detection of Foodborne Pathogens

Table 33.3 β-Lactamases Identified in Gram-Negative Bacilli β-Lactamase

Gene

Type

Note

TEM SHV OXA

blaTEM blaSHV

Not yet released ten types; withdrawn two types; repeatedly named five types Not yet released 17 types; withdrawn one types; repeatedly named five types

blaOXA

1-161 1-105 1-127

CTX-M CMY

blaCTX-M

1-69

Repeatedly named two types

blaCMY

1-36

IMP

blaIMP

1-23

VIM

blaVIM

1-18

KPC GES

blaKPC

1-4

blaGES

1-9

PER

blaPER

1-3

VEB

blaVES

1-6

SME

blaSME

1-3

KPC-1 is identical with KPC-2

Source: Based on Lahey Clinic Website, http://www.lahey.org/studies/webt.asp. Last modified, January 7, 2008.

a potential of dissemination is present, the distribution of bacteria harboring blaSME remains somewhat limited because the gene is encoded on the chromosome. This contrasts with IMP-1, a plasmid-encoded metallo-β-lactamase, identified first in a clinical isolate of S. marcescens in Japan in 1994.35 Shortly afterward, the blaIMP-1 gene was identified worldwide from a variety of clinical isolates, such as Pseudomonas aeruginosa, Klebsiella pneumoniae, Escherichia coli, and Acinetobacter baumannii.36–38 To date, 23 different types of IMP β-lactamase have been identified in S. marcescens.17 Another type of metallo-β-lactamase, VIM-2, was also found in S. marcescens in 2002.39 Table  33.4 summarizes all 44 types of β-lactamases identified in S. marcescens,32–35,39–67 33.1.3.2  The Outer Membrane Barrier The outer membrane of Gram-negative bacteria serves as a diffusion barrier to extracellular solutes including antibiotics. Porins in the outer membrane function as transmembrane channels and can be divided functionally into two classes: nonspecific diffusion channels allowing the energy­independent passage of some solutes; and substrate-specific diffusion channels.68,69 The channels limit the penetrations of large or hydrophobic compounds, thereby contributing to intrinsic resistance against antimicrobial agents. OmpC (40 kDa) and OmpF (41 kDa) are two porins in S. marcescens characterized by Hutsul’s group in 1997. They are nonspecific channels involved in β-lactam permeation, and their amino acid sequences are 71% and 68% homologous with E. coli OmpC and OmpF.70,71 Alteration of their expression is associated with decreased outer membrane permeability, and defects can induce β-lactam resistance.72,73

33.1.4 Mechanisms of S. marcescens Resistance to Fluoroquinolones Fluoroquinolones (e.g., norfloxacin, ciprofloxacin, ofloxacin, and enoxacin) are the first entirely man-made

antibiotics, and are one of the first choice agents against S. marcescens. In Gram-negative bacteria, they selectively inhibit DNA gyrase, thus blocking DNA replication and transcription. Three resistance mechanisms have been described in S. marcescens: first, gyrase mutations which decrease affinity to fluoroquinolones; second, reduced intracellular drug accumulation due to active efflux; and third, plasmid-mediated qnr proteins which bind to DNA gyrase and protect it. 33.1.4.1  Mutations of gyrA DNA gyrase is composed of subunits A and B encoded by the gyrA and gyrB genes. Mutations of gyrA are associated with fluoroquinolone resistance.74 Mutations conferring highlevel resistance have been mapped to a small region of the gene, designated as the quinolone resistance-determining region (QRDR). These mutations most frequently occur in the Ser-83 and Asp-87 codons, and have been confirmed in 60 clinical isolates of Enterobacteriaceae, including S. marcescens.75 Compared to wild type S. marcescens gyrA sequence (GenBank accession No. AF052260), six quinolone-resistant clinical isolates of S. marcescens showed single amino acid substitutions at Gly-81, Ser-83 or Asp-87, indicating that a point mutation at the QRDR of gyrA confers resistance to fluoroquinolones (Table 33.5).75,76 33.1.4.2 Active Efflux Pump Some bacteria can pump toxic substances and antibiotics outside the cells through proteinaceous transporters, called active efflux pumps. Six major superfamilies of these transporters have been described: ATP-binding cassette (ABC), major facilitator superfamily (MFS), small multidrug resistance, (SMR), multidrug endosomal transporter (MET), resistance nodulation division (RND), and multi antimicrobial resistance (MAR).77 Of these, the ABC pumps are primary trans­ porters utilizing ATP hydrolysis as a source of energy, while the rest are secondary transporters using a proton or sodium

463

Serratia

Table 33.4 β-Lactamases Produced by Serratia spp. β-Lactamase TEM-1 TEM-2 TEM-3 (CTX-1) TEM-24 (CAZ-6) TEM-24b TEM-47 TEM-52 TEM-AQ SHV-2 SHV-5 SHV-9 (SHV-5a) SHV-12 (SHV-5-2a) CTX-M-1 CTX-M-2 CTX-M-3 CTX-M-9 CTX-M-14 CTX-M-15 (UOE-1) SME-1 SME-2 SME-3 SFC-1 KPC-2 GES-1 SFO-1 FONA-3 BES-1 CKH-1 OHIO-1 IMP-1 IMP-6 IMP-10 IMP-11 VIM-2 SFH-I AmpC AmpC-like SRT-1 SRT-2 OXA-1 OXA-1 like OXA-10 OXA-30 OXA-31

Ambler’s Classification

Serratia Species and Strain

GenBank Accession No.

Reference

A A A A A A A A A A A A A A A A A A A A A A A A A A A A A B B B B B B C C C C D D D D D

marcescens ES-71 marcescens marcescens CF-15 marcescens marcescens marcescens marcescens marcescens S5 marcescens marcescens marcescens marcescens marcescens marcescens marcescens KHM604 marcescens liquefaciens SUN-188 marcescens BB 1758 marcescens S6 marcescens 4126 marcescens OC7554 fonticola UTAD54 marcescens marcescens fonticola CUV fonticola SF13 marcescens Rio-5 marcescens 42039 marcescens 076 marcescens TN9106 marcescens KU3838 marcescens S16 marcescens 1297 marcescens YMCU1591 fonticola UTAD54 marcescens SR50 marcescens SC8247 marcescens GN16694 marcescens marcescens 3105 marcescens marcescens SCH88050909 marcescens marcescens 466

AY538702 X54606* X64523* X65253* – – Y13612* X97254 EU376967* EU441171* X98105* EU137700* AM003904* EF592571* AB185839 AY092058* AF462398 EU118602* Z28968 AF275256 AY584237 AY354402 AF481906* DQ333893* AB003148* AJ251241 AF234999 – M33655* S71932 AB040994 AB074433* AB074437* AY030343 AF197943 X52964 – AB008454 AY524276 J02967* AY008291* AF453998 EU339234* EF067840

40 41 42 43 44 45 46 47 48 45 49 46 40 50 46 40 – 51 32 33 34 52 53 54 55 – 56 57 58 35 59 113 60 39 61 62 63 64 65 60 46 66 67 –

* Registered under other Gram-negative bacilli.

gradient as a source of energy. Unlike the classical enzymesubstrate or ligand-receptor recognization, an efflux system is able to recognize a number of compounds depending on their physico-chemical properties, such as hydrophobicity, aromaticity and ionizable characters. Most antibiotics are amphiphilic molecules possessing both hydrophilic and

hydrophobic properties. Therefore, they can be recognized and pumped out by many efflux systems. Table 33.6 summarizes the recently described efflux pumps that make S. marcescens resistant to fluoroquinolone. The genes are mostly chromosomal, and their expression can be induced and regulated by antibiotics.78–83

464

Molecular Detection of Foodborne Pathogens

Table 33.5 Mutations of gyrA Associated with Fluoroquinolone Resistance in S. marcescens Amino Acid and Codon at Position

MIC (μg/ml) Strain Wild-type ATCC 13880

CPFX 0.5

OFLX 1

81 Gly (GGT)

83 Ser (AGC)

1221

4

8

Cys (TGT)

–

9745

2

4

5591

8

 ≥ 16

1568

 ≥ 16

 ≥ 16

4

 ≥ 16

3.2

25.6

– – – – –

87 Asp (GAC)

Reference 75

Mutant

1969 DSWH 101

75

Arg (CGC)

– – – –

–

Asn (AAC)

75

Tyr (TAC)

76

Ile (ATC) Arg (AGA)

75 75 75

Notes: –, no change versus the sequence of ATCC 13880; CPFX, ciprofloxacin; OFLX, ofloxacin.

Table 33.6 Multidrug Efflux Pumps Identified in S. marcescens Pump SmdAB SdeXY SmfY SdeAB SsmE

Gene

Superfamily

Substrate

Strain

smdA, smdB sdeX, sdeY smfY sdeA, sdeB ssmE

ABC RND MFS RND SMR

NFLX, CPFS, TC, TPPCl NFLX, EM, TC, EB, ACR NFLX, OFLX, ACR, TPPCl, EB CPFS, NFLX, OFLX, CP, EB CPFS, NFLX, ACR, EB

NUSM8906 NUSM8906 NUSM8903 ATCC13880 NUSM8903

GenBank Accession No. AB360548 AB104882 AB251607 AY168756 AB361065

Reference 79 80 81 82 83

Notes: ACR, acriflavine; CP, chloramphenicol; CPFS, ciprofloxacin; EB, ethidium bromide; EM, erythromycin; NFLX, norfloxacin; OFLX, ofloxacin; TC, tetracycline; TPPCl, tetraphenyl-phosphonium chloride.

33.1.4.3  Qnr Determinants QnrA, the first plasmid-mediated resistance determinant to fluoroquinolone, was identified in a K. pneumoniae strain in the United States in 1998.84 Qnr proteins may directly protect DNA gyrase from quinolones, leading to the increase of minimum inhibitory concentrations (MICs).85 QnrA, QnrB, and QnrS have now been confirmed in various Enterobacteria.86–88 Among 166 isolates of S. marcescens, 0.6%, 1.2%, and 0.6% were positive for qnrA1, qnrB1, and qnrB4, respectively.89 The qnrS1 gene is also found in S. marcescens.90

In Gram-negative bacteria, the main resistance mechanism is production of aminoglycoside-modifying enzymes.91,92 The enzymes are classified as aminoglycoside N-acetyltransferase (AAC), aminoglycoside O-adenylyltransferase (ANT, also called aminoglycoside nucleotidyltransferase), and aminoglycoside O-phosphotransferase (APH).93 These enzymes are also sub-classified depending on the site of modification and unique resistance profiles. About 50 aminoglycoside-modifying enzymes have been identified, mostly from Gram-negative bacteria.94 Table 33.7 summarizes aminoglycoside-modifying enzymes identified in S. marcescens.

33.1.5 Mechanisms of S. marcescens Resistance to Aminoglycosides

33.1.5.1 Aminoglycoside N-acetyltransferases (AAC) AAC are classified into four classes by whether they modify a 1-, 3-, 6'-, or 2'-amino group. AAC(6') is widely distributed in Gram-negative bacteria, and some of these enzymes can modify clinically important aminoglycosides. AAC(6')-I acetylates amikacin, tobramycin, and netilmicin, but not gentamicin; while AAC(6')-II acetylates gentamicin, tobramycin, and netilmicin, but not amikacin.94–97 The aac(6')-Ia, -Ib, and -Ic have been identified in Serratia spp.98–100 and aac(6')-Il was also found in S. marcescens isolate YMC/KSM51 (GenBank Accession No. AY884051). The aac(6')-Ic gene has been detected in all

Aminoglycosides, such as gentamicin, tobramycin, amikacin, and streptomycin, are commonly used in the clinical setting, and gentamicin is recommended as the second choice agent against S. marcescens.16 An aminoglycoside is a glycoside with an amino substituent. Aminoglycosides interfere with protein synthesis by binding to the A-site of the 30S subunit of bacterial ribosome, thus disturbing elongation of the peptide chain, and causing misreading of mRNA.

465

Serratia

Table 33.7 Aminoglycoside-Modifying Enzymes Identified in S. marcescens Enzyme AAC(6')-Ia AAC(6’)-Ib AAC(6’)-Ic AAC(6’)-Il AAC(6’)-II AAC(3)-I AAC(3)-II ANT(2’’)-Ia ANT(3’’)-Ia ANT(4')-II APH(3’)-Ia APH(3’)-Ib ANT(3'')-Ii/AAC(6')-IId

Gene aac(6’)-Ia aac(6’)-Ib aac(6’)-Ic aac(6')-Il aac(6’)-II aac(3)-Ia aac(3)-IIa ant(2'')-Ia ant(3'')-Ia ant(4')-IIa aph(3')-Ia aph(3')-Ib ant(3’’)-Ii/aac(6’)-IId

Alterative Name aacA1 aacA4 – aacA7 – aacC1 aacC2 aadB aadA1 – aphA1 aphA2 –

Substrate

Strain or Plasmid

GenBank Accession No.

Reference

AMK, TOB AMK, TOB AMK, TOB AMK, TOB GM, TOB GM GM, TOB TOB, GM SM, SPCM AMK, TOB KM KM SM, SPCM, KM

– pAZ007 SM16 YMC/KSM51 823 pUO901 01 SCH88050909 SCH88050909 – – – SCH88050909

M18967* M23634 M94066 AY884051 AY675225* S68049 AY663802* AF453999 AF453999 M98270* V00359* M20305* AF453998

98 99 100 – 97 102 103 66 66 105 98 98 66

Notes: AMK, amikacin; GM, gentamicin; KM, kanamycin; SM, streptomycin; SPCM, spectinomycin; TOB, tobramycin. * Registered under other Gram-negative bacilli.

S. marcescens strains studied (n = 351), but not in other species (n = 3,842).100,101 Therefore, aac(6')-Ic is considered as a molecular marker for S. marcescens. 3-N-acetyltransferase AAC(3), is also widely distributed in Gram-negative bacteria and can acetylate gentamicin.97,102,103 A DNA probe for aacC1 was used to test 21 isolates of gentamicin-resistant S. marcescens at the Seattle Veterans Administration Medical Center, and only two isolates carried the gene.97 In Belgium, however, the aacC1 gene was detected frequently in Serratia spp. and 87% of aminoglycoside-resistant strains (n = 123) carried the gene encoding AAC(3)-Ia.98 33.1.5.2 Aminoglycoside O-adenylyltransferases (ANT) The 2"-O-adenylyltransferase, ANT(2"), is responsible for low-level resistance to gentamicin, tobramycin, and kanamycin.104 About 80% (n = 83) and > 50% (n = 101) of aminoglycoside-resistant Serratia spp. were positive for ANT(2")-Ia and ANT(3")-Ia, respectively, as confirmed by DNA hybridization.98 Recently, the genes for ANT(2")-Ia and ANT(3")-Ia were identified in a conjugative plasmid from S. marcescens SCH88050909, a multi-resistant clinical isolate.66 ANT (4')-II was also detected in S. marcescens.105 33.1.5.3 Aminoglycoside O-phosphotransferases (APH) The 3'-O-phosphotransferase, APH(3'), is the most important because it is frequently identified in clinic, and mediates resistance to kanamycin and neomycin. At least seven subclasses of the APH(3') enzymes have been defined in ­Gram-negative bacilli.94 DNA probes for APH-(3')I and APH(3')-II were used to screen for aph genes in Gram-negative bacilli.106 More than 50% of aminoglycoside-resistant Serratia spp. (n = 238) were positive for aph(3')-Ia.98

33.1.5.4 Bifunctional Aminoglycoside Resistance Enzyme Some enzymes exhibit two distinct activities, that is, they are bifunctional. The fusion of two genes forms a two-domain protein with two active sites. The first bifunctional aminoglycoside resistance enzyme, AAC(6')/APH(2''), was discovered in Enterococcus faecalis in 1986.107 Another enzyme, ANT(3'')-Ii/AAC(6')-IId, has recently been discovered from S. marcescens. The enzyme catalyzes adenylation and acetylation of aminoglycoside, and its gene is located on a plasmid.66,108 Although bifunctional enzymes are rare in bacteria, their emergence may become a new clinical threat.

33.1.6 Laboratory Identification of S. marcescens As a member of the Enterobacteriaceae family, S. marcescens is readily cultured and identified by traditional methods in clinical laboratories and can be distinguished from other bacteria in the family by several properties, such as the production of DNase, lipase, gelatinase and red pigment (prodigiosin); casein hydrolysis; tryptophan- and citratedegradation; negative for methyl red and indol tests; positive for lactic acid production; motile with peritrichous flagella.3 Production of red pigment is a well-known property of S. marcescens, however, less than 10% of its strains actually produce the pigment.4 Molecular approaches are also reported to identify S. marcescens as shown below.

33.2  Methods A number of molecular approaches are available for identification of bacteria in the Enterobacteriaceae family such as Escherichia, Klebsiella, and Serratia. One approach

466

utilizes PCR to amplify 16S rRNA gene followed by restriction enzyme digestion or nucleotide sequencing analysis,109,110 which can be relatively time-consuming. Another approach employs primers and probe targeting 16S rRNA gene in a real-time PCR detection format, thus permitting rapid result turnover.111,112 We present below two protocols for identification of S. marcescens, one of which is a PCR-restriction digestion technique described by Lu et al.109 and the other is a real-time PCR procedure reported by Iwaya et al.111

33.2.1  PCR and Restriction Enzyme Digestion109 Sample preparation. Approximately 105 CFU of bacteria are washed in 1 ml of a lysis buffer (1% Triton X-100, 10 mM Tris pH 8.0, and 1 mM EDTA) by centrifugation at 13,000 × g for 5 min. The pellet is resuspended in 100 μl of the same lysis buffer and then boiled in a water bath for 30 min to release the DNA. After removing the cell debris by centrifugation at 13,000 × g for 5 min, the supernatant is saved for PCR. To identify S. marcescens in cerebrospinal fluid specimens, 500 μl of cerebrospinal fluid is centrifuged at 13,000 × g for 5 min. The pellet is resuspended in 180 μl of sterile distilled water, and the DNA is then isolated with the QIAamp Tissue Kit. PCR amplification. PCR mixture (50 μl) ­containing about 50 ng of template DNA, 1 × PCR buffer, a 0.2 μM concentration of each PCR primer (U1 and U2), a 0.2 mM concentration of each deoxynucleoside triphosphate, and 2.5 U of Taq DNA polymerase in a total volume of 50 μl is prepared. The nucleotide sequence of primer U1 is 5′-CCAGCAGCCGCGGTAATACG-3′ (correlating to nt 518–537 of the E. coli 16S rRNA gene, and the nucleotide sequence of primer U2 is 5′-ATCGG(C/T) TACCTTGTTACGACTTC-3′, correlating to nt 1513–1491 of the same gene. PCR amplification is undertaken in a thermal cycler with the following program: one cycle of 94°C for 10 min; 35 cycles of 94°C for 1 min, 55°C for 1 min, and 72°C for 2 min; one cycle of 72°C for 10 min. Upon completion, 5 μl of PCR product is electrophoresed on a 1% agarose gel to determine the size of the product, which should be 996 bp from the members of Enterobacteriaceae family. Restriction endonuclease digestion. 5 μl of each PCR product is digested with restriction enzyme HaeIII in a total volume of 20 μl. After incubation for 2 h at the recommended temperature, the digested DNA is electrophoresed on a 6% polyacrylamide gel to confirm the presence of 217, 210, 180, 161, 126, 68, and 34 bp fragments (which are shared by Enterobacter cloacae, E. coli, K. pneumoniae, and S. marcescens but not by other members of Enterobacteriaceae family). Then, 5 μl of each PCR product is digested with restriction enzyme DdeI in a total volume of 20 μl to produce two fragments of 757 and 239 bp for E. coli and K. pneumoniae and three fragments of 474, 283, and 239 bp for S. marcescens and E. cloacae. Next, 5 μl of each PCR product is digested with restriction enzyme BstBI in a total volume of 20 μl to separate E. coli and S.

Molecular Detection of Foodborne Pathogens

marcescens (both of which are undigested by BstBI) from K. pneumoniae, and E. cloacae (both of which are digested into two fragments of 876 and 120 bp).109

33.2.2 Real-Time PCR111 Sample preparation. S. marcescens isolates are grown in LB broth (Difco Laboratories, Detroit, MI) or nutrient agar (Eiken Chemical, Tokyo) at 37°C. Bacterial DNA is extracted from 200 μl of the suspension containing 1 × 107 CFU/ml with the QIAamp DNA mini kit (Qiagen, Tokyo) following the manufacturer’s instructions. The DNA is dissolved in 200 μl of Tris–hydrochloride pH 8.0, and 20 μl aliquots of the DNA solution are used for the real-time PCR assay. Real-time PCR amplification and detection. For the real-time PCR detection of S. marcescens, the primers (SMSF and SMSR) and probe (SMSP) based on the 16S rRNA gene sequence of S. marcescens are used: SMSF, 5′-GGTGA GCTTAATACGTTCATCAATTG (nt 435–460); SSR, 5′-G CAGTTCCCAGGTTGAGCC (nt 595–613); SMSP, 5′-TGCGC TTTACGCCCAGTAATTCCGA (nt 534–558). The primers SMSF and SMSR enable amplification of a fragment of 179 bp. The probe SMSP contains the reporter dye FAM (6-carboxyfluorescein; Applied Biosystems, Foster City, CA) at the 5′ end and the quencher dye TAMRA (6-carboxytetramethylrhodamine; Applied Biosystems) at the 3′ end. The PCR ­mixture (50 μl) consists of 400 nM of the primers, 80 nM of the probe, 25 μl of Taqman universal prepared mixture and 20 μl of the template. The real-time PCR is performed with an ABI 7900HT sequence detector (Applied Biosystems), with the following cycling programs: one cycle of 95°C for 10 min (to activate AmpliTaq Gold) and 50 cycles of 95°C for 15 sec and 60°C for 1 min. The intensity of fluorescence (ΔRn) is calculated by subtracting the baseline fluorescence from the actual fluorescence signal data. The threshold cycle (CT) is defined as the cycle number at which the reporter fluorescence exceeded the threshold value, a parameter defined as 10 SD above the baseline fluorescence. This real-time PCR assay allows rapid and specific detection of S. marcescens septicemia in 2.5 h and with 200 μl of blood.

33.3 Conclusions and Future Perspectives S. marcescens is a well known cause of nosocomial infections in humans. It is ubiquitous in the environment, can contain genes encoding many types of resistance to antibiotics, and can spread this resistance to other Gram-negative bacilli. Therefore, molecular detection and surveillance of the resistant genes harbored by S. marcescens becomes more and more important, to assess and control their spread and colonization in hospitals, and to guide treatment of infections.

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Serratia













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34 Shigella*

Benjamin R. Warren ConAgra Foods

Keith A. Lampel

Food and Drug Administration

Keith R. Schneider University of Florida

Contents 34.1 Introduction.................................................................................................................................................................... 471 34.1.1 Morphology and Classification ........................................................................................................................ 471 34.1.2 Biology, Epidemiology, and Pathogenesis ....................................................................................................... 472 34.1.3 Current Technologies for Diagnosis of Shigella............................................................................................... 475 34.2 Methods.......................................................................................................................................................................... 478 34.2.1 Reagents, Supplies, and Equipment.................................................................................................................. 478 34.2.2 Sample Preparation, Enrichment, and Bacteriological Culture........................................................................ 478 34.2.3 Nested PCR Detection of Shigella.................................................................................................................... 479 34.3 Conclusions and Further Perspectives ........................................................................................................................... 480 References.................................................................................................................................................................................. 481

34.1  Introduction 34.1.1 Morphology and Classification Morphology and classification. Shigella spp. are the causative agents of shigellosis, or “bacillary dysentery.” Shigella spp. are Gram-negative, facultatively anaerobic, nonsporeforming, nonmotile rods. Typically, Shigella spp. do not ferment lactose, are lysine-decarboxylase, acetate, and mucate negative and do not produce gas from glucose, although some six S. flexneri serotypes have been reported to produce gas.1,2 Shigellae belong to the bacterial family Enterobacteriaceae, and are almost genetically identical to Escherichia coli and are also closely related to Salmonella and Citrobacter spp.3 There are very few biochemical or genetic characteristics that can distinguish Shigella spp. from enteroinvasive E. coli (EIEC).4 Based on biotyping, the genus Shigella is divided into four serogroups: S. dysenteriae (serogroup A; 15 serotypes), S. flexneri (serogroup B; eight serotypes divided into 11 subserotypes), S. boydii (serogroup C; 20 serotypes), and S. sonnei (serogroup D; one serotype).5 Molecular evolutionary relationship between Shigella and EIEC. It is generally accepted that Shigella are within the species E. coli. Recent studies have indicated that

Shigella, like the other forms of pathogenic E. coli, derived from multiple evolutionary origins, suggesting convergent evolution of the Shigella phenotype.6 Ribotyping of 75 strains of Shigella, 13 strains of EIEC and 72 E. coli strains from the E. coli Reference (ECOR) Collection resulted in S. sonnei, S. flexneri and most S. dysenteriae ribotypes being closely related to phylogenic group D, while S. dysenteriae serotype 1 strains were closely related to phylogenic group B1 and S.  boydii strains were spread between phylogenic group  D and B1.7 In contrast, the ribotypes of EIEC strains were widely distributed among phylogenic groups A, B1, and B2. This evidence suggests that Shigella and EIEC derived from different origins. Pupo et al.6 sequenced eight housekeeping genes from four regions of the chomosome for 46 strains of Shigella representing all four serotypes. Three distinct clusters of Shigella were identified and although S. sonnei and S. dysenteriae serotype 1, 8, and 10 did not group in the main three clusters, they fell well within the species E. coli.6 As with the study by Rolland et al.7 S. boydii serotype 13 was distantly related to the other Shigella strains. Cluster 1 contained most of the S. boydii and S. dysenteriae strains along with S. flexneri serotypes 6 and 6A. Cluster 2 contained seven S. boydii strains and S. dysenteriae serotype 2. Cluster 3 contained

* A portion of the material presented in this chapter has been previously published by Taylor and Francis (www.informaworld.com). Warren, B.R. Parish, M.E. and Schneider, K.R. Crit. Rev. Food. Sci. Nutr. 46, 551–567, 2006.

471

472

S. flexneri serotypes 1–5 and S. boydii serotype 12. Unlike the results from ribotyping, the use of multiple genes for phylogenic analysis revealed greater genetic diversity among the strains of Shigella, further suggesting that Shigella derived from different evolutionary origins. Fukiya et al.8 used comparative genomic hybridization microarray analysis to compare the gene content of E. coli K-12 with that of 22 pathogenic E. coli and Shigella strains. When compared to the E. coli K-12 genome, the genomes of S. sonnei, S. boydii and S. flexneri 2a were missing only 613, 533 base pairs and 716 open reading frames (ORFs). The genomes of the other pathogenic E. coli strains were missing similar numbers of ORFs. Subsequent phylogenic analysis revealed a close relationship between thee of four EIEC strains and the three strains of Shigella, which suggests EIEC and Shigella form a single E. coli pathovar.8,9 The complete genomes of S. boydii serotype 4 (strain 227), S. dysenteriae serotype 1 (strain 197) and S. sonnei (strain 046) have recently been sequenced.9 Comparative genomics among the newly sequenced Shigella genomes and the previously sequenced genomes of S. flexneri 2a (strain 301) and E. coli K-12 (strain MG1655) supported previous work by Fukiya et al.8 While the genomes of Shigella share most of their genes with that of E. coli K-12, the Shigella phenotype is a result of the gain and loss of functions through bacteriophage-mediated gene acquisition, insertion sequence (IS)-mediated DNA rearrangements and formation of pseudogenes.9 For example, the chomosome and virulence plasmid (VP) of S. sonnei strain 046 contained 327 and 28 intact IS elements and 67 and 68 partial IS elements, respectively. In contrast, the E. coli K-12 genome contained only 37 intact IS elements and seven partial IS elements. In an attempt to resolve conflicting theories on the molecular evolution of Shigella, Yang et al. constructed tree sets of phylogenic trees: a chomosomal tree consisting of 23 housekeeping genes from 46 strains of Shigella and EIEC, a chomosomal tree using four housekeeping genes from 19 E. coli strains from the ECOR Collection and 46 strains of Shigella/ EIEC, and a VP tree using five genes outside of the ­cell-entry region from 38 Shigella/EIEC strains. Both chomosomal trees grouped Shigella into three main clusters (C1, C2, and C3) with five outliers. In the chomosomal trees, clusters C2 and C3 were more closely related; however, in the VP tree clusters C1 and C2 were more closely related. This data led the authors to conclude that diverse ancestral VPs entered diverse origins of E. coli to result in the various Shigella species that share pathogenic properties.

34.1.2  Biology, Epidemiology, and Pathogenesis Survival. There is limited data on the prevalence of Shigella among food handlers or on food products. One group of investigators found Shigella in the stools of four out of 283 examined food handlers in Irbid, Jordan.10 Mensah et al.11 evaluated 511 food items from the streets of Accra, Ghana, from which S. sonnei was isolated from one sample of

Molecular Detection of Foodborne Pathogens

macaroni. It was noted that the macaroni was served using bare hands instead of clean utensils, which may have led to the S. sonnei contamination. Wood et al.12 examined foods from Mexican homes, commercial sources in Guadalajara, Mexico, and from restaurants in Houston, TX, for contamination with bacterial enteropathogens. While no Shigella was isolated from foods sampled from 12 Houston restaurants or from food commercially prepared in Guadalajara, Mexico, four isolates were obtained from meals prepared in Mexican homes. These studies demonstrate the importance of proper food handling and the role food handlers in the transmission of Shigella. The U.S. Food and Drug Administration (FDA) has also investigated the presence of foodborne pathogens, including Shigella, in imported and domestic produce.13–15 From imported produce, Shigella was found on three of 151 cantaloupe samples, two of 84 celery samples, one of 116 lettuce samples, one of 84 parsley samples, and two of 180 scallion samples.13 From domestically grown produce, Shigella was found on one of 164 cantaloupe samples, three of 93 scallion samples, and one of 90 parsley samples.15 Shigella spp. are generally heat sensitive,16 acid resistant,17 salt tolerant,18 and can withstand low levels of organic acids.19 Freezing (–20°C) and refrigeration (4°C) of contaminated foods typically improves survival of Shigella, however growth is not observed at these temperatures. Shigella can survive in water with little decline in population20 and on fruits and vegetables, often beyond the product’s shelf-life.20–22 Produce packaged under vacuum or modified atmosphere have also supported the survival of Shigella.23 Low pH foods (vegetable juices and prepared salads) can support survival of Shigella when held at refrigerated temperatures.20,24 Many cooked foods (when recontaminated post-cook) support the survival of Shigella and growth may be observed under temperature abuse situations.25 In addition to foods, inanimate objects (fomites) can serve as vectors for transmission of Shigella and there have been several reports on the survival of Shigella on various types of surfaces.26–28 Outbreaks and common foods. Historically, Shigella has been thought of as a waterborne pathogen29 and outbreaks have been commonly associated with contaminated community water sources, such as swimming pools, that were un- or under-chlorinated.30,31 Foodborne outbreaks of Shigella are also common, and are usually associated with foods that are subjected to processing/preparation by hand, are exposed to a limited heat treatment, or are served/delivered raw to the consumer.22 Many of the foods associated with Shigella outbreaks are considered ready-to-eat (RTE) foods, meaning there is no application of heat (or other kill step) prior to the point of consumption. Various types of produce and/ or products that contain produce items, such as prepared salads, have been implicated in foodborne outbreaks of Shigella (Table 34.1). Community gatherings, such as church picnics, where food items have been prepared in large portions and potentially been subjected to temperature abuse have also been associated with outbreaks of Shigella.

473

Shigella

The FDA has investigated the presence of foodborne pathogens, including Shigella, on imported and domestic produce.13–15 In fiscal year (FY) 1999, the FDA initiated a survey of high-volume imported produce items.13 In the FY 1999 survey, Shigella was isolated from nine of 671 samples: three of 151 cantaloupe samples, two of 84 celery samples, one of 116 lettuce samples, one of 84 parsley samples, and two of 180 scallion samples.13 In 2000, the FDA also initiated a survey focused on high-volume domestic fresh produce.15 In the domestic survey, Shigella was isolated in five of 665 samples: one of 164 cantaloupe samples, three of 93 scallion samples, and one of 90 parsley samples.15 A 300-sample follow up assignment on imported fresh produce was assigned in FY 2001; however, Shigella was not isolated from any of the 257 samples in that survey. Epidemiology. The four serogroups of Shigella differ in epidemiology.32 S. dysenteriae is primarily associated with epidemics32 with serotype 1 associated with the highest fatality rate (5–15%).33 S. flexneri predominates in areas of endemic infection, while S. sonnei has been implicated in source outbreaks in developed countries.34 S. boydii has been associated with source outbreaks in Central and South America but is most commonly restricted to the Indian subcontinent. S. boydii is rarely isolated in North America; however slight increases in the numbers of isolates have been observed in both 2003 and 2004.35 According to the CDC Emerging Infections Program, Foodborne Diseases Active Surveillance Network (FoodNet), Shigella was the third most reported foodborne bacterial pathogen in 2005.36 Of 17,252 laboratory-diagnosed cases, Shigella accounted for 2,736 cases (15.9% of total cases) behind only Salmonella (6,655 cases) and Campylobacter (5,712 cases).36 From 1996–1998 to 2006 the estimated

Table 34.1 Selected Foodborne Outbreaks Associated with Shigella Year

Serogroup

Food Product(s) Implicated

annual incidence of Shigella spp. in the U.S. decreased by 35%.36 S. sonnei continued to be the most isolated serogroup in the U.S. followed by S. flexneri, S. boydii and S. dysenteriae, respectively (Table 34.2). FoodNet data on shigellosis in the U.S. collected from 1996 to 1999 were analyzed for trends in demographic variability.37 The overall incidence of shigellosis was highest among the following groups: children aged 1–4 years, male patients, African Americans, Hispanics, and Native Americans.37 There were also marked demographic differences between infection with S. sonnei and S. flexneri with respect to age, sex, and race. While the incidence of both S. sonnei and S. flexneri were higher among those aged 1–4 years, there was a second peak of S. flexneri infection among those aged 30–39 years.37 The incidence of S. sonnei among men and women were similar; however the incidence of S. flexneri among men was almost twice that of women.37 In addition, the incidence of S. sonnei among African Americans and whites was higher than that of S. flexneri, while the incidence of S. flexneri was higher among Native Americans.37 Pathogenesis. Invasion of epithelial cells by shigellae involves four steps: entry into epithelial cells, intracellular multiplication, intra- and intercellular spreading, and killing of the host cell.38 The invasion process is controlled by a ­plasmid-encoded genes. The plasmid contains invasion plasmid antigen (ipa) genes which encode four highly immunogenic polypeptides; IpaA, IpaB, IpaC, and IpaD. Studies in which Tn5 insertions affecting the expression of the ipa genes reveal that expression of ipaB, ipaC, and ipaD is strongly associated with entry, while an ipaA mutant is invasive but at reduced ability as compared to wild type.39 Invasion is also mediated by the virF gene, which is located on the vp, and the virR gene, which is located on the chomosome. The virF gene encodes a 30-kDa protein that positively regulates the expression of the ipa genes and a plasmid gene icsA (also known as virG), which encodes intra- and intercellular spread. Environmental factors which affect the expression of virF are not known. The virR gene is a repressor of the plasmid invasion genes in a temperature-dependant manner.38 When shigellae are grown at 30°C they do not express any of the Ipa polypeptides and are therefore noninvasive, however, shigellae grown at 37°C are fully invasive and all plasmid polypeptides are empressed.38

1983 1986

S. sonnei S. sonnei

Tossed salad Shedded lettuce

1986

S. sonnei

Raw oysters

1987

S. sonnei

Watermelon

1988

S. sonnei

Uncooked tofu salad

1989

S. flexneri 4a

German potato salad

1992

S. flexneri 2

Tossed salad

1994 1995–1996

S. sonnei S. sonnei

Iceberg lettuce Fresh pasteurized milk cheese

1996

S. flexneri

Salad vegetables

Table 34.2 Percentage of Shigella Isolates in the United States Reported by PHLIS in Recent Years

1998

S. sonnei

Uncooked, chopped curly parsley

Serogroup

2002

2003

2004

2005

1998

S. flexneri

Restaurant-associated, source unknown

S. sonnei

83.5%

80.2%

68.9%

74.4%

1999

S. boydii 18

Bean salad (parsley or cilantro)

S. flexneri

12.2%

14.4%

17.2%

13.6%

2000

S. sonnei

Five layer bean dip

S. boydii

0.8%

1.1%

1.8%

1.2%

2001

S. sonnei

Raw oysters

S. dysenteriae

0.3%

0.4%

0.4%

0.5%

2002

Shigella spp.

Greek-style pasta salad

2004

S. sonnei

Raw carrots

Ungrouped Total isolates

3.2% 12,992

3.9% 11,552

11.7% 9,343

10.3% 10,484

474

Once ingested, Shigellae move through the gastrointestinal tract to the colon, where they translocate the epithelial barrier via M cells that overlay the solitary lymphoid nodules.40 Upon reaching the underlying M cells, Shigella infects the macrophages and induces cell death.40 Infected macrophages release interleukin-1β, which elicits a strong inflammatory response.41 Once released from the macrophage, Shigella will enter the epithelial cells, also called enterocytes, which predominately line the colon via membrane ruffling and macropinocytosis. Epithelial cells produce inflammatory cytokines in response to bacterial invasion, therefore increasing inflammation of the colon.40 Macropinocytosis involves the cell extending its membrane as formations known as pseudopodia, which will engulf large volumes and close around them forming a vacuole. In order for macropinocytosis to occur, actin must be polymerized and myosin, an actin binding protein, must be present.42 Common stimuli that induce macropinocytosis include cytokines and bacterial antigens. Shigellae immediately disrupt phagocytic vacuoles allowing entry into the host cell cytoplasm. Once in the cytoplasm, Shigellae multiply rapidly. Sansonetti et al.43 observed the generation time for S. flexneri in HeLa cells to be approximately 40 min. Sustaining efficient intracellular growth requires the acquisition of host cell nutrients. Although production of Shiga toxin (ST) facilitates the availability of host cell nutrients, no relation between its production and intracellular growth rates can be observed.44 Likewise, no relation between Shiga-like toxin (SLT) production and intracellular growth rate can be observed.43,45 Since little free iron exists within mammalian host cells, Shigella spp. must also express high-affinity iron acquisition systems. In order to obtain iron, Shigella spp. synthesize and transport the siderophores aerobactin and enterobactin46 and utilize a receptor/transport system in which iron is obtained from heme. Siderophores (aerobactin and enterobactin) are low molecular weight iron binding compounds that remove iron from host proteins. Enterobactin is produced by some but not all Shigella spp.47 while aerobactin is synthesized by S. flexneri and S. boydii48 and some S. sonnei.49 Headley et al.50 demonstrated that aerobactin systems, although active in extracellular environments, are not expressed intracellularly. This suggests that siderophore-independent iron acquisition systems can provide essential iron during intracellular multiplication.50 The capacity for Shigella to spread intracellularly and infect adjacent cells is critical in the infection process.38 Intraand intercellular spreading is controlled by the icsA (virG) gene located on the vp. The icsA gene encodes the protein IcsA, which enables actin-based motility51 and intercellular spread.52 IcsA is a surface-exposed outer membrane protein consisting of thee distinctive domains: a 52 amino acid N-terminal signal sequence, a 706 amino acid α-domain, and a 344 amino acid C-terminal β-core.53–55 The α-domain is the exposed portion and the β-core is embedded in the outer membrane.55

Molecular Detection of Foodborne Pathogens

IcsA is distributed at one pole of the outer membrane surface. This asymmetrical distribution allows the polar formation of actin tails, and thus polar movement of Shigella within host cell cytoplasm. The polar localization of IcsA is primarily affected by two events: (i) the rate of diffusion of outer membrane IcsA;56–59 and (ii) the specific cleavage of IcsA by the protease IcsP (SopA).60–62 Rate of diffusion of outer membrane IcsA is directly affected by the O side chains of the membrane lipopolysaccharide (LPS). Sandlin et al.57 demonstrated this relationship with a S. flexneri LPS mutant, BS520 that does not make any O-antigen. As compared with a wild-type strain of S. flexneri, which polymerized actin at one pole, the LPS mutant strain polymerized actin in a nonpolar fashion. Expression of LPS that does not have any O side chains causes an even distribution of IcsA over the entire outer membrane.57,63 Composition of the C-terminal one-third of the IcsA α-domain is also required for polar localization as well as polar movement of S. flexneri.55 A S. flexneri mutant, in which a segment of this section was deleted, was unable to polymerize actin in a polar fashion or move unidirectionally.64 The icsP gene encodes the outer membrane protease IcsP (also called SopA) that cleaves laterally diffused IcsA, thus promoting polar localization.61 An E. coli K-12 strain, engineered to express the icsA gene, was shown to diffuse IcsA along its outer membrane.63 When the same E. coli K-12 strain was engineered to express the icsP gene with the icsA gene, the number of bacteria which polymerized actin at one pole increased.63 The N-terminal two-thirds of the IcsA α domain is essential for mediating actin assembly in Shigella host cells.40 This portion of IcsA contains six glycine rich repeats which interact with the Wiskott–Aldrich syndrome protein (N-WASP). N-WASP is composed of distinct domains: PH, a pleckstrin homology domain; IQ, a calmodulin binding domain; GBD, a GTPase binding domain that binds Cdc42; PRR, a proline-rich region; V, a G-actin-binding veroprolin homology domain; C, cofilin homology domain; A, a C-terminal acidic amino acid segment.40,65 The VCA domain of N-WASP activates and interacts with the Arp2/3 complex. The IcsA–NWASP–Arp2/3 complex mediates rapid actin filament growth at the barbed end, including cross-linking between the elongated actin filaments.40 The resulting network of actin filaments allows Shigella to gain a propulsive force with which to move in the cytoplasm of host cells.40 Cdc42 (an N-WASP activator), profilin (an actin monomer-binding protein), and cofilin (which depolymerizes actin) also have roles in efficient actin assembly. Cdc42 binds to the GBD domain of N-WASP preventing the intramolecular interaction between the C-terminal acidic amino acids and the basic amino acids of the GBD, thereby forcing the N-WASP complex to unfold to its activated form.40 Cdc42 independent activation of the N-WASP complex by IcsA has been reported for actin-based motility in Shigella, however efficient entry into cells was reported as Cdc42 dependent.66 The role of Cdc42 in invasion and motility is still somewhat controversial. Profilin delivers monomeric

475

Shigella

actin to sites of actin assembly.67 Although profilin has been shown not to be absolutely essential for actin-based motility, it is required for maximum rates of movement.68 Cofilin generates actin monomers from the filamentous actin. By disassembling unneeded actin filaments within the tail, cofilin might work to free up actin for incorporation into newly generated filaments.67 Intercellular spreading is dependent upon an actin-based motility mechanism.63 Shigella cells first form a membrane bound protrusion into an adjacent cell. This protrusion must distend two membranes: one from the donor cell, and another from the recipient cell.69 As the protrusion pushes further into the recipient cell, it is taken up by the recipient cell resulting in the bacteria enclosed in a double-membrane vacuole.63 Intercellular spread is completed when Shigella rapidly escapes from the double-membrane vacuole, releasing it into the cytosol of the secondary cell. Monack and Theriot63 observed intercellular spread of an E. coli K-12 strain expressing the icsA and icsP genes in HeLa cells. As expected, they found the E. coli K-12 strain spread to adjacent cells and enclosed in a double-membrane vacuole as well as free in the cytosol of the adjacent cells. The early killing of host cells by Shigella is mediated by the vp. Sansonetti38 demonstrated the inability of noninvasive S. flexneri to kill host cells whereas the invasive species killed efficiently and rapidly. Noninvasive strains can not kill host cells, since this requires that the Shigella be intracellular. Early killing of host cells involves metabolic events that rapidly drop the intracellular concentration of ATP, increase pyruvate, and arrest lactate production.70 S. dysenteriae type 1 strains produce a potent toxin known as Shiga toxin (STX). Interestingly, STX, a potent cytotoxin produced by S. dysenteriae serotype 1, does not play a role in the early killing of host cells. Fontaine et al.44 constructed a Tox– mutant strain of S. dysenteriae serotype 1 and found that the mutant killed as efficiently as the wild-type strain. Although the toxin is not necessary to sustain an infection, its expression increases the severity of disease. Three biologic activities associated with STX are cytotoxicity, enterotoxicity, and neurotoxicity, while the one known biochemical effect is the inhibition of protein synthesis.71 STX is considered the prototype to a family of toxins known as SLT, which are similar in structure and function, and share the same receptor sites. Perhaps the most widely known human pathogen that produces SLT’s is E. coli O157:H7, which has two toxin variants (SLT I and SLT II). STX is composed of two polypeptides: an A subunit (32,225 MW) and five B subunits (7,691 MW each).71 The B subunits mediate binding to cell surface receptors, which have been identified as glycolipids containing terminal galactose-α(1-4)galactose disaccharides such as galabiosylceramide and globotriasylceramide (Gb3).72,73 The A subunit, once inside the host cell cytoplasm, acts enzymatically to cleave the N-glycosidic bond of adenine at nucleotide position 4324 in the 28S rRNA of the 60S ribosomal unit.71 Two enterotoxins, Shigella enterotoxin 1 (SHET 1) and Shigella enterotoxin 2 (SHET 2), have been characterized

and are believed to play a role in the clinical manifestation of shigellosis.74 SHET 1, which is chomosomally encoded, was only prevalent in isolates of S. flexneri 2a. SHET 2, however, is encoded on the large vp, and was detected in all Shigella isolates tested except several isolates that had lost their plasmid. Medical importance. Worldwide, shigellosis is estimated to cause 600,000 deaths annually and is one of the most common causes of infectious diarrhea in both developed and developing countries.75 In the U.S. approximately 14,000 cases of shigellosis are reported each year; however the actual number of cases may be as high as 20 times greater since milder cases are often not diagnosed or reported.76 In 2006, the incidence of shigellosis in the U.S. was reported to be 6.09 per 100,000 persons.36 The infective dose for Shigella is very low: ten cells of S. dysenteriae and 500 cells of S. sonnei.77 Populations such as the very young, the very old, or persons with decreased immune function may be at more risk for infection. Due to the low infective dose of Shigella, person-to-person transmission is common. Typical symptoms of infection include bloody diarrhea, abdominal pain, fever, and malaise. Although the mechanism is unknown, seizures have been reported in 5.4% of shigellosis cases involving children.78 Chonic sequelae from S. dysenteriae serotype 1 infections can include hemolytic uremic syndrome (HUS), while S. flexneri infections are associated with later development of reactive arthritis, especially in persons with the genetic marker HLA-B27.76 Reactive arthritis is characterized by joint pain, eye irritation, and painful urination.76 Treatment of shigellosis includes both rehydration and antimicrobial therapy. The administration of antibiotics serves to alleviate dysenteric syndromes such as fever and abdominal cramps, and reduces the average duration of pathogen excretion and the risk of transmission and/or lethal complications.79 Unfortunately, resistance to multiple antibiotics among Shigella strains has progressively increased.80 Among Shigella isolates from five Canadian provinces (1997–2000), high rates of resistance to ampicillin (65%) and trimethoprim-sulfamethoxazole (TMP-SMX) (70%) were observed.81 Similarly, Shigella isolates in the U.S. (1999–2002) were resistant to ampicillin (78%) and TMP-SMX (46%).82 In the U.S. Shigella isolates remain susceptible to ciprofloxacin and ceftriaxone.82 The WHO has recommended ciprofloxacin as the first-line antibiotic for treatment of shigellosis, although fluoroquinolone-resistant Shigella spp. have been documented in several countries.83

34.1.3 Current Technologies for Diagnosis of Shigella Conventional detection of Shigella in food. Traditional microbiological techniques make use of selective and differential media for the enrichment and isolation of Shigella. Many variants of enrichment and plating media have been investigated for optimal recovery, often with conflicting results. Although Shigella is readily isolated from clinical

476

samples, food samples are more problematic. Isolation of Shigella from food samples can be inhibited by indigenous microflora, especially the coliform bacteria and Proteus spp.2 The addition of the antibiotic novobiocin to liquid and solid media has been shown to improve the isolation of S. flexneri and S. sonnei from investigated foods.2 Contamination of food products with Shigella results primarily through a food handler with poor personal hygiene; therefore the concentration of Shigella may be very low compared to that of the indigenous microflora.84 Currently, selective media are not available that will adequately suppress the growth of background microflora; therefore Shigella is often overgrown by competing microorganisms.84 More research is needed to determine more appropriate selective media and enrichment conditions for the isolation of Shigella from food samples. Evaluations of the FDA BAM Shigella culture method have produced favorable results for prepared salads and fruits/vegetables, and highly variable results for raw products, such as ground beef and oysters.85– 87 Immunological methods, such as latex agglutination (LA), enzyme immunoassay (EIA), or immunomagnetic separation (IMS), have limited application for the detection of Shigella in foods. LA requires the prior isolation of a suspect colony on solid media; therefore, LA serotyping kits for Shigella that are commercially available cannot be used for detection of Shigella directly from a food sample, but rather can confirm or aid in characterization of suspected Shigella colonies. Commercially available EIA test kits for the detection of Shigella are seldom used for the routine analysis of foods in the U.S. IMS techniques have been reported in literature to concentrate Shigella from clinical samples for downstream detection processes such as EIA88 or PCR89–91 but no IMS kits are known to be commercially available at time of this publication. IMS has been shown to eliminate PCR inhibitors inherent to fecal samples, and IMS-PCR methods reported for the diagnosis of shigellosis in children with severe diarrhea were more than two times as effective then the conventional culture method.90 These studies have significance with respect to Shigella detection in foods, however further investigations are needed to evaluate the suitability of the IMS techniques for use with food samples. Virulent S. sonnei produce smooth colonies, termed form I, which result from expression of the O-antigen. Unlike other Shigella species, the LPS genes (rfc and rfb) of S. sonnei are located on the large vp, which can be spontaneously lost at high frequency. Sansonetti et al.92 investigated the stability of form I plasmids and observed 1–45% plasmid loss from re-streaking form I colonies onto MacConkey agar and incubating 24 h at 37°C. When the large vp is lost, rough (avirulent) colonies, termed form II, are produced that express the Enterobacteriaceae R1 LPS core.92,93 A defective mutant of form II S. sonnei LPS, termed R-form, is characterized by an incomplete core region.93 As antibodies specific for S. sonnei O-antigen will bind form I, but not form II or R-form LPS, immunological detection methods with specificity for the O-antigen of Shigella are compromised when the vp is lost.

Molecular Detection of Foodborne Pathogens

Molecular detection of Shigella in foods. For the detection of Shigella, the integration of conventional bacteriological methods and DNA-based technology has been used successfully for the resolution of at least one major foodborne outbreak in the U.S. In 2000, a multistate outbreak was reported to the CDC that was epidemiologically associated with eating a five-layered bean dip.94 A total of 406 cases were identified in ten U.S. states. The etiological agent was identified from the analysis of patient’s stool specimens to be S. sonnei. Initial attempts to isolate S. sonnei from samples of the five-layered bean dip using conventional bacteriological methods were unsuccessful. Different layers of the bean dip were separated and subjected to enrichment followed by concurrent analysis by PCR and bacteriological culture (enrichment and bacteriological culture in accordance with the Shigella culture method of the FDA, Bacteriological Analytical Manual).95 Four subunits of the cheese layer from the bean dip tested positive for Shigella by PCR and eventually S. sonnei was isolated from two of these subunits.96 Pulse-field gel electrophoresis (PFGE) confirmed that the isolates from clinical specimens matched those isolated from the bean dip. The use of PCR allowed investigators to identify the contaminated component of the five-layer bean dip and facilitated the isolation of the outbreak strain. PCR methods for detection of Shigella in food have previously demonstrated higher sensitivity than comparable culture methods.87 PCR assays for Shigella spp. have targeted the invasion associated locus (ial),88,97 the virA gene,98,99 or the ipaH gene87,100–103 for amplification. Due to their genetic similarity, EIEC are detected by all reported PCR methods targeting the ial, virA or ipaH genes. The ial and virA genes are located on the large vp (sometimes referred to as the invasion plasmid), however sequencing of the S. flexneri genome104 revealed the ipaH gene to be encoded multiple times on both the chomosome and the large vp. Thus, detection of the ipaH gene is possible in the event the large vp has been lost, which may occur when food samples are stored for a prolonged periods of time prior to analysis.103 PCR methods have been previously reported for the detection of Shigella for in mussels,99 mayonnaise,98 lettuce, shimp, milk, and blue cheese,97 environmental water samples,102 and fresh tomatoes.87,105 Table 34.3 summarizes PCR and other rapid methods for the detection of Shigella in foods and feces. Immunocapture universal primer PCR (iUPPCR) has been recently reported.106 Universal primer PCR employs primers designed against highly conserved regions, such as 16S rRNA genes, thus the resulting PCR can be used for amplification of almost any bacteria. Typically, the sequence of the resulting amplicon would be further analyzed for identification of the bacteria. In iUPPCR however, immunocapture is employed for specificity and universal primer PCR is used for detection of captured bacteria. Monoclonal antibodies specific for individual serotypes of S. dysenteriae, S. flexneri, S. boydii, or S. sonnei were coated into the wells of a 96-well polystyrene plate. Cross-reactivity tests demonstrated the specificity of the monoclonal antibodies to the strain level.

477

Shigella

Table 34.3 Selected Rapid Methods for Detection of Shigella Serogroup

Target

Matrix

Sample Preparation

Detection

Sensitivity

S. sonnei S. boydii S. flexneri

ipaH

Tomatoes

FTA filtration

Nested PCR

<101 cfu/tomato

ipaH

Water

SE, boiled extract

Semi-nested PCR

≥1.1 × 101 cfu/ml

S. dysenteriae 1

virA

Mayonnaise

CEP

PCR

102–103 cfu/ml

S. dysenteriae

virA

Mussels

PE, CEP

PCR

101–102 cfu/ml

S. flexneri

plasmid DNA

Lettuce

Alkaline denaturation

PCR

1.0 × 104 cfu/g

S. flexneri

ial

Various foods

SE, BDC

Nested PCR

1.0 × 101 cfu/g

LPS, ial

Feces

IMS

PCR

1.0-1.5 × 101 cfu/ml

LPS

Rectal swabs

PE, DPSM

EIA

ND

LPS

Various foods

None

Biosensor

≥4.9 × 104 cfu/ml

LPS, ial

Feces

IMS

PCR

ND

LPS, ial

Feces

IMS

PCR

1.0 × 101 cfu/g

LPS, 16S rRNA

Sewage

Immunocapture

UPPCR

5.0 × 101 cfu/ml

LPS

Feces

IMS

EIA

1.0 × 103 cfu

S. sonnei form I S. dysenteriae S. flexneri S. boydii S. sonnei S. dysenteriae S. dysenteriae 1 S. flexneri S. sonnei S. dysenteriae 1 S. flexneri S. dysenteriae S. flexneri S. boydii S. sonnei S. dysenteriae 1 S. flexneri

Abbreviations: SE, selective enrichment; PE, pre-enrichment; ND, not determined; LPS, lipopolysaccharide; EIA, enzyme immunoassay; BDC, buoyant density centrifugation; UPPCR, universal primer PCR; DPSM, direct plate on selective media; CEP, chemical extraction/ethanol precipitation.

Wells were challenged with bacteria and captured bacteria were detected using universal primer PCR, i.e., PCR with primers specific for 16S rRNA sequences conserved among closely related bacteria. The detection limit for shigellae in broth was 5.0 × 102 cfu/ml.106 Presumably, detection of up to 96 pathogens sharing the conserved 16S rRNA sequence could be accomplished in the same plate using specific monoclonal antibodies for bacterial capture (differentiation of cell types) and the universal primer PCR for detection. Further investigations of this method with respect to food samples as opposed to bacteria in broth are be required. Advances in oligonucleotide microarrays have further enabled the simultaneous detection and identification of bacterial foodborne pathogens. Oligonucleotide microarrays involve the ordered immobilization of specific probes to a solid surface followed by hybridization with labeled sample DNA. After hybridization, development of the label can allow identification/enumeration of complementary sample DNA sequences. Kakinuma et al.107 describes an oligonucleotide microarray using probes specific for the gyrB gene to detect and differentiate E. coli, Shigella, and Salmonella. PCR amplified regions of the gyrB gene were fluorescently labeled and hybridized to detection probes immobilized on glass slides. Based on reaction patterns, three species of Shigella, seven serovars of Salmonella, and one strain of E. coli were

correctly identified at the species level. Identification at the subspecies level of Salmonella was problematic when multiple serovars were present in the same sample due to the overlap of microarray patterns. Assay formats such as this could be expanded to potentially detect and identify a wide range of enteric pathogens. Repetitive sequence-based PCR (rep-PCR) has been demonstrated for the identification and molecular typing of members of the family Enterobacteriaceae, including Shigella.108,109 In rep-PCR, primers are designed complementary to bacterial interspersed repetitive sequences and the regions lying outward of the primers are amplified resulting in DNA fragments of varying lengths. These fragments can then be separated by electrophoresis to form a bacterial fingerprint unique to individual bacterial strains. The DiversiLabTM System (Spectral Genomics, Inc. Houston, TX) generates such fragments by rep-PCR that are then analyzed using microfuidics lab-on-a-chip and the Agilent 2100 Bioanalyzer (Agilent Technologies, Inc. Palo Alto, CA). Results can be visualized in several formats including dendograms, electropherograms, gel-like image, or scatter plots.109 Rep-PCR technology has been shown to be reproducible and can allow the differentiation of species, subspecies and strains in as little as 4 h. Some Shigella serogroups show > 95% similarity with the DiversiLabTM

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Enteric Kit Beta version, however the DiversiLabTM Shigella kit offers greater differentiation among serogroups and strains.109 Strains of E. coli (nonpathogenic, enterohemorragic, and enterotoxigenic) were effectively differentiated from Shigella spp. using rep-PCR,108,109 however no enteroinvasive E. coli strains were included in the studies.

34.2  Methods In an ideal situation, the analysis of foods for the presence of Shigella spp. should include a rapid means for detection, as well as isolation of the pathogen from food samples. Molecularbased methods can have a significant impact on the analysis of foods, particularly as a screening tool. Since many PCR-based assays are not able to discriminate live versus dead cells, the application of such techniques should address this concern. The nested PCR assay for the detection of Shigella from foods presented below was designed to demonstrate that increased amplification is due to growth of live cells. The objective of using the nested PCR assay is to confirm that 620 bp DNA fragments amplified is the primary PCR were from Shigella. Primers used in the nested PCR are directed to internal sequences within the 620 bp amplicon targeted in the primary PCR. In the method presented below, samples are processed at 0, 6, and 24 h for PCR analysis, and bacteriological analysis is performed at 6 and 24 h. In essence, the bacteriological analysis and PCR are performed concurrently. PCR analyses are performed at the 0 and 6 h time points (as a screen) and at 24 h and as a confirmation of suspect colonies on agar plates. A strengthening of the band in the 6 h sample over the 0 h sample is suggestive of growth and may provide evidence that detection was a result of the presence of live cells if cultural isolation of the pathogen is unsuccessful.

34.2.1 Reagents, Supplies, and Equipment For preparation of media and reagents, refer to the FDA Bacteriological Analytical Manual110 (Table 34.4). The number in parentheses following each medium refers to its listing in the media section of the manual.

34.2.2 Sample Preparation, Enrichment, and Bacteriological Culture (i) Preparation of 0 h PCR analyte: Individual samples are placed into a sterile Whirl-Pak bag. For the weight of the product, add an equal volume of BPB (e.g., if one sample weighs 250 g, add 250 ml BPB for a 1:1 dilution). Gently shake the bag with contents for 5 min using an orbital shaker at 100 rpm. From this rinse, place 2 × 10 ml into two 15-ml centrifuge tubes. Centrifuge the tubes at 6,000 rpm for 5 min. Remove supernatant (to avoid leaving behind any inhibitory compounds, aspiration may be a better alternative than decanting the supernatant). Suspend/combine the pellet from each tube in 1.0 ml of PBS and transfer the suspended material to a 1.5-ml

Molecular Detection of Foodborne Pathogens

microcentrifuge tube. For PCR template, see preparation protocol below. Note: If the 1:1 dilution contains large amounts of plant material and debris, decant approximately 2 × 10 ml of the liquid rinse into two 15-ml centrifuge tubes. Perform a slowspeed spin (2,000 rpm for 2 min) to remove debris. Carefully transfer 2 × 10 ml of the supernatants to fresh 15 ml centrifuge tubes and continue with protocol above at the centrifuge step. (ii) Sample enrichment: Transfer the remaining sample rinse from above (e.g., 105 ml if the original sample was 250 g) to a sterile 500 ml Erlenmeyer flask containing an equal volume of 2 × Shigella broth with novobiocin (final concentration of novobiocin in the enrichment broth should be 0.5 µg/ml; therefore, in the 2 × recipe, the concentration of novobiocin is 1.0 µg/ml). Adjust pH, if necessary, to 7.0 ± 0.2 with sterile 1 N NaOH or 1 N HCl. Incubate at 40°C ± 2°C for 6 h, aerobic conditions, with shaking (125–150 rpm). (iii) Preparation of 6 h PCR analyte and bacteriological analysis: After 6 h of enrichment, transfer 2 × 10 ml into two 15-ml centrifuge tubes. Centrifuge the tubes at 6,000 rpm for 5 min. Remove supernatant (to avoid leaving behind any inhibitory compounds, aspiration may be a better alternative than decanting the supernatant). Suspend/combine the pellet from each tube in 1.2 ml of PBS and transfer the suspended material to a 1.5 ml microcentrifuge tube. Spread two 100-µl aliquots evenly over two Shigella Rainbow agar plates. Incubate plates overnight (16–24 h) at 37°C ± 2°C. The remaining 1.0 ml of suspended material is the 6 h PCR analyte. Store at 4°C if not processed immediately. Continue incubation of the remaining Shigella broth enrichment (approx. 190 ml if original sample was 250 g) at 40°C ± 2°C for 16 h, aerobic conditions, with shaking (125– 150 rpm). (iv) Preparation of 24 h PCR analyte and bacteriological analysis: From the enriched Shigella broth (total enrichment time of 24 h from steps ii and iii above), transfer 2 × 10 ml into two 15-ml centrifuge tubes. Centrifuge the tubes at 6,000 rpm for 5 min. Remove supernatant (to avoid leaving behind any inhibitory compounds, aspiration may be a better alternative than decanting the supernatant). Suspend/combine the pellet from each tube in 1.0 ml of PBS and transfer the suspended material to a 1.5-ml microcentrifuge tube. Store at 4°C if not processed immediately. Also from the enriched Shigella broth, streak a 10-µl loopful onto Shigella Rainbow, HE, and XLD agar plates immediately after enrichment. Incubate plates overnight (16–24 h) at 37°C ± 2°C. (v) Identification of suspect colonies: The characteristics of Shigella colonies on selective agar plates are: (1) Shigella Rainbow agar: Shigella colonies are mauve (a dull pink-purple color). The color is darkest in

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Shigella

Table 34.4 List of Media, Reagents, Supplies and Equipment Required Microbiological Media

Reagents

Supplies and Equipment

Shigella broth with novobiocin (M136) Shigella Rainbow agar (BioLog) Hektoen Enteric agar (M61)

1N Sodium hydroxide solution (R73)

Pipette tips, DNase-free, aerosal barrier (10 µl, 100 µl, 100 µl)

1N Hydrochloric acid (R36) Phosphate buffered saline (PBS) (11.42 g NaCl, 1.08 g Na2HPO4, 0.315 g KH2PO4, 1.5 L dH2O pH 7.4)

Whirl-Pak bags, sterile Centrifuge tubes, sterile, 15 ml

Xylose Lysine Desoxycholate agar (M179) Tryptic Soy Agar (TSA), prepared as slants Butterfield’s phosphate buffer (BPB)

0.5 × Tris-acetate EDTA (TE) buffer pH 8.3

Microcentrifuge tubes, DNase-free, sterile (1.5 ml, 0.2 ml)

Distilled water, sterile (dH2O)

Centrifuge w/ rotor sized for 15 ml and 1.5 ml tubes

100 bp DNA ladder as molecular weight marker PCR primers

Orbital shaker Erlenmeyer flask, sterile, 500 ml

PCR master mix, containing buffer, Taq DNA polymerase, dNTPs, MgCl2 (HotStart TaqTM Master Mix from Qiagen) Agarose gel, 1%, molten, w/ 5 ppm ethidium bromide Gel loading dye

Incubator, shaking (125–150 rpm), 40°C ± 2°C

Disposable bacteriological loops, sterile, 10 µl Incubator, 37°C ± 2°C Heat block w/ blocks sized for 1.5 ml tubes Electrophoresis gel tray Thermocycler S. flexneri strain 2457M -ATCC 29903™ Disposable plate spreaders, sterile UV transilluminator

the center of the colony and fades to near colorless around the edges. (2) HE agar: Shigella colonies are approx. 2–3 mm and appear green. (3) XLD agar: Shigella colonies are 1–2 mm diameter, transparent or red, smooth colonies. (vi) Preparation of PCR template from suspect colonies: From six to 12 well-isolated suspect colonies, pick part (not the entire colony) of each colony and inoculate to TSA agar slants. The locations of selected colonies should be clearly marked using a permanent marker on plates since they will need to be further analyzed below. Incubate slants at 37°C ± 2°C for 20 h. Incubated slants are stored at 4°C in case further biochemical characterization of an isolate is required. From the remaining portions of the selected colonies, transfer a small amount of colony material to a 1.5-ml microcentrifuge containing 150 µl of dH2O. Heat at 100°C for 5 min, cool on ice and then use 1.0 µl for PCR analysis as described in step i below. (vii) Preparation of PCR template from sample enrichments: Centrifuge the 1.0 ml suspended cells (prepared as 0 h, 6 h, and 24 h PCR analytes in steps i, iii, and iv above) at 6,000

rpm for 5 min to pellet bacterial cells. Remove as much of the supernatant as possible (to avoid leaving behind any inhibitory compounds; aspiration may be a better alternative than decanting the supernatant). Suspend cells in 50–500 µl of 1 × PBS (this will depend upon the size of the pellet). If the pellet is hardly visible, then add the smallest amount of 1 × PBS. Heat the cell suspension at 100°C for 5 min, cool on ice and centrifuge at 14,000 rpm for 3 min. Transferring the supernatant to another tube is not necessary; use the supernatant as template without disturbing the pellet. The PCR templates from the 0 and 6 h enrichments may be held at 4°C overnight to be processed by PCR at the same time as the 24 h template.

34.2.3 Nested PCR Detection of Shigella (i) PCR primer: Prepare primer solutions at 10 pmol/µl and store at –20°C. The sequences of the primers are as follows: (a) ipaHF: 5´-GTT CCT TGA CCG CCT TTC CGA TAC CGT C-3´ (b) ipaHR: 5´-GCC GGT CAG CCA CCC TCT GAG AGT AC-3´

480



(c) (d) (e) (f)

Molecular Detection of Foodborne Pathogens

607: 5´-ATGCTTGATGTTAAAAATACAGGAG-3´ 608: 5´-GCTCTTTCTTGTATGCACTTGTG-3´ ipaH3: 5´-CCA CTG AGA GCT GTG AGG-3´ ipaH4: 5´-TGT CAC TCC CGA CAC GCC-3´

(ii) Primary PCR: Prepare 25 µl reaction mixtures for each of the DNA templates (0 h, 6 h and 24 h) consisting of 12.5 µl 10 × PCR master mix, 2.5 µl primer ipaHF, 2.5 µl primer ipaH, DNA template (2.5 µl for sample enrichment templates and 1 µl for colony templates) and dH2O. The thermocycling conditions are: one cycle of 95°C for 15 min; 30 cycles of 94°C for 1 min, 60°C for 1 min, and 72°C for 1 min; indefinite hold at 4°C. Note: It is recommended that the positive control strain (S. flexneri 2457M) be run in parallel with test samples to ensure that the reagents are working properly. Positive control template should be prepared as described in step vi above. To make sure positive test samples were not due to cross contamination, the primary PCR may be repeated by substituting the control primers 607 and 608 for primers ipaHF and ipaH in the PCR mix described above. For PCR amplification using the control primers 607 and 608, cycling conditions are as described above except the annealing temperature is 56°C. Amplification (PCR) with primers 607 and 608 will yield a 1.8-kb amplicon from the control strain (S. flexneri 2457M) only; amplification from all other Shigella spp. will yield a 1.0-kb amplicon. Primers 607 and 608 are specific for the mxiC gene of Shigella, therefore if no PCR product is observed when these primers are used the sample should not be interpreted as being negative for Shigella; the vp could be completely or partially deleted in isolates from food samples. Alternatively, S. flexneri 2457M can be differentiated from other Shigella isolates by its resistance to the antibiotic kanamycin­ (50 µg/ml). (iii) Electrophoresis of PCR product: Transfer 10 µl of the PCR products to a clean, sterile 0.2 ml microcentrifuge tube containing 2 µl of tracking dye and mix using a vortex. If needed, a quick spin on a centrifuge may be used to return all of the mixture to the bottom of the tube. Load the 12 µl mixture into a single well of a 1% agarose gel submerged in TE buffer in an electrophoretic gel tray. A 100-bp DNA ladder should be loaded in at least one of the outside wells. Separate the bands by electrophoresis and visualize bands using a UV transilluminator. Note: Do not add dye directly to the PCR product. Hold the remaining PCR product at 4°C or store at –20°C. A 620-bp product is expected from positive samples in the primary PCR. If a 620-bp amplicon is seen on the agarose gel, proceed to the nested PCR procedure for confirmation. Nested PCR Prepare 25 µl reaction mixtures consisting of 12.5 µl 10 × PCR master mix, 2.5 µl primer ipaH3, 2.5 µl primer ipaH4, 1.0 µl DNA template (product from primary PCR in step i above) and 6.5 µl dH2O. The thermocycling conditions are the same as described in step i above.

Notes: Usually if 1.0 µl of the primary PCR product (620 bp fragments) is used directly as template in the nested PCR, then three bands may be visible on the gel; the correct band at 290 bp, and two larger bands at approx. 400 and 500 bp. The two larger bands are due to primers ipaHF and ipaH being carried over from the primary PCR reaction. The larger amplified bands are generated from the combinations of ipaHF-ipaH4 and ipaH-ipaH3. To avoid these extra bands the primary PCR product may be diluted 1:10 or 1:100 in dH2O and 1.0 µl of the diluted primary PCR product may be used as template in the nested PCR. In some cases, faint bands around 400 and 500 bp may be observed even when a 1:10 dilution of the primary PCR product is used as template in the nested PCR. A band at 290 bp is confirmation for the presence of Shigella. Interpretation and reporting of results Since this method includes PCR analysis of both sample enrichments and suspect colonies, careful interpretation of results is required. The following rules should be used for the interpretation of data and reporting of results. (1) The presence of Shigella cannot be ruled out if any of the primary PCR analyses of sample enrichments are positive, AND/OR the primary PCR analysis of any suspect colony is positive. (2) The presence of Shigella is confirmed positive if any of the nested PCR analyses of sample enrichments are positive, AND/OR the nested PCR analysis of any suspect colony is positive. (3) The sample is negative for Shigella if both the primary PCR analyses of sample enrichments are negative AND microbiology with primary PCR analyses of suspect colonies are negative.

34.3 Conclusions and Further Perspectives Molecular methods, such as PCR, for the detection of Shigella in food have been effectively used by regulatory agencies for the investigation of foodborne outbreaks. Although there are clear advantages (e.g., detection limit and time to results) for the use of molecular methods over conventional culture methods, there are still challenges and limitations associated with molecular methods. Virtually all molecular methods for the detection of Shigella in foods require the use of an enrichment step for sufficient sensitivity (as is true for other bacterial pathogens as well). As discussed above, selective media for the enrichment of Shigella are not currently available that can adequately suppress the growth of background microflora; therefore, the sensitivity of current molecular detection is dependant on the ability of Shigella grow in the presence of potential food matrix and/or competing microflora inhibition. Another limitation of PCR methods with respect to the detection of Shigella in foods is the potential detection of dead cells. Since bacterial DNA can remain intact in a food

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Shigella

sample for an extended period of time following cell death, the results of PCR methods in the absence of culture isolation have been challenged to have resulted from the detection of dead cells. Given that the reservoir for Shigella is limited to humans and high primates, the presence of viable Shigella cells and/or Shigella DNA in a food sample suggests the food was prepared/produced in unhygienic conditions and may not be fit for human consumption. To help address the issue of dead versus live cell detection, the PCR method provided in this chapter was designed to support detection of viable Shigella in the event of unsuccessful culture isolation. The strengthening of PCR signal (e.g., band intensity) as enrichment time increases suggests the multiplication of Shigella, thereby providing evidence that positive detection resulted from viable cells. Most current PCR assays for the detection of Shigella in food target invasion-related genes. Since these invasion­related genes are shared among Shigella spp. and EIEC, both pathogens, if present at sufficient populations in a sample, would be detected. The PCR method provided in this chapter will detect both Shigella and EIEC. While this may be viewed as a limitation of the method, EIEC and Shigella spp. form a single pathovar and the presence of either pathogen in a food product suggests that the food was prepared/produced in unhygienic conditions. Although EIEC is an uncommon foodborne etiological agent, foodborne outbreaks have occurred. The symptoms of shigellosis and EIEC infections are virtually identical, therefore methods used for diagnosis of disease and subsequent investigations may benefit from simultaneous detection of Shigella and EIEC. Future research on molecular methods for the detection of Shigella (and other foodborne bacterial pathogens) in foods should focus on the development of universal sample preparation techniques. Current sample preparation techniques are not applicable to all food matrices due mainly to chemical/ physical differences among samples (e.g., viscosity, presence of inhibitory substances, etc.). With the diversity of foods for which microbiological analyses are required, a universal sample preparation technique would vastly improve the utility of molecular methods. In an ideal world, a universal sample preparation technique could be applied to a food sample, without need for enrichment, and molecular methods, such PCR, could provide same-day, sensitive and specific detection of bacterial pathogens.

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482 23. Satchell, F.B. et al. The survival of Shigella sonnei in shedded cabbage. J. Food Prot., 53, 558, 1990. 24. Bagamboula, C., Uyttendaele, M., and Debevere, J. Acid tolerance of Shigella sonnei and Shigella flexneri. J. Appl. Microbiol., 93, 479, 2002. 25. Islam, M.S., Hasan, M.K., and Khan, S.I. Growth and survival of Shigella flexneri in common Bangladeshi foods under various conditions of time and temperature. Appl. Environ. Microbiol., 59, 652, 1993. 26. Spicer, C.C. The survival of Shigella sonnei on cotton theads. J. Hyg., 57, 210, 1959. 27. Nakamura, M. The survival of Shigella sonnei on cotton, glass, wool, paper and metal at various temperatures. J. Hyg., 60, 35, 1962. 28. Islam, M.S. et al. Survival of Shigella dysenteriae type 1 on fomites. J. Health Popul. Nutr., 19, 177, 2001. 29. Smith, J.L. Shigella as a foodborne pathogen. J. Food Prot., 50, 788, 1987. 30. Fleming, C.A. et al. An outbreak of Shigella sonnei associated with a recreational spray fountain. Am. J. Public Health, 90, 1641, 2000. 31. Centers for Disease Control and Prevention (CDC). Shigellosis outbreak associated with an unchlorinated fill-and-drain wading poolóIowa, 2001. Morb. Mortal. Wkly. Rep., 50, 797, 2001. 32. Ingersoll, M. Groisman, E.A., and Zychlinsky, A. Pathogenicity islands of Shigella. Curr. Topics Microbiol. Immunol., 264, 49, 2002. 33. Centers for Disease Control and Prevention (CDC). PHLIS surveillance data. Shigella: annual summary, 2002. Available at: http://www.cdc.gov/ncidod/dbmd/phlisdata/shigella.htm. 2003. Accessed 05 Nov 2004. 34. Hale, T. Genetic basis of virulence in Shigella species. Microbiol. Rev. 55, 206, 1991. 35. Centers for Disease Control and Prevention (CDC). Preliminary FoodNet data on the incidence of infection with pathogens transmitted commonly through food—10 States, United States, 2005. Morb. Mortal. Wkly. Rep., 55, 392, 2006. 36. Centers for Disease Control and Prevention (CDC). Preliminary FoodNet Data on the Incidence of Infection with Pathogens transmitted commonly through food—10 States, 2006. Morb. Mortal. Wkly. Rep., 56, 336, 2007. 37. Shiferaw, B. et al. Trends in population-based active surveillance for shigellosis and demographic variability in FoodNet sites, 1996–1999. Clin. Infect. Dis, 38 (Supp13), S175–80, 2004. 38. Sansonetti, P.J. Genetic and molecular basis of epithelial cell invasion by Shigella species. Rev. Infect. Dis., 13(Suppl 4), S285, 1991. 39. Tran Van Nhieu, G., Ben-Ze’ev, A., and Sansonetti, P.J. Modulation of bacterial entry into epithelial cells by association between vinculin and the Shigella IpaA invasion. EMBO J., 16, 2717, 1997. 40. Suzuki, T., and Sasakawa, C. Molecular basis of the intracellular spreading of Shigella. Infect. Immun., 69, 5959, 2001. 41. Zychlinsky, A. et al. Interleukin 1 is released by murine macrophages during apoptosis induced by Shigella flexneri. J. Clin. Invest., 94, 1328, 1994. 42. Stendahl, O.I. et al. Distribution of actin-binding protein and myosin in macrophages during spreading and phagocytosis. J. Cell. Biol., 84, 215, 1980. 43. Sansonetti, P.J. et al. Multiplication of Shigella flexneri within HeLa cells: lysis of the phagocytic vacoule and plasmid-mediated contact hemolysis. Infect. Immun., 54, 461, 1986.

Molecular Detection of Foodborne Pathogens 44. Fontaine, A., Arondale, J., and Sansonetti, P.J. Role of shiga toxin in the pathogenesis of bacillary dysentery studied using a tox- mutant of Shigella dysenteriae 1. Infect. Immun., 56, 3099, 1988. 45. Clerc, P.L. et al. Plasmid-mediated early killing of eucaryotic cells by Shigella flexneri as studied by infection of J774 macrophages. Infect Immun., 55, 521, 1987. 46. Vokes, S.A. et al. The aerobactin iron transport system genes in Shigella flexneri are present within a pathogenicity island. Mol. Microbiol., 33, 63, 1999. 47. Perry, R.D., and San Clemente, C.L. Siderophore synthesis in Klebsiella pneumoniae and Shigella sonnei during iron deficiency. J. Bacteriol., 140, 1129, 1979. 48 Lawlor, K.M., and Payne, S.M. Aerobactin genes in Shigella spp. J. Bacteriol., 160, 266, 1984. 49. Payne, S.M. Iron and virulence in the family Enterobacteriaceae. CRC Crit. Rev., Microbiol. 16, 81, 1988. 50. Headley, V. et al. Expression of aerobactin genes by Shigella flexneri during extracellular and intracellular growth. Infect. Immun., 65, 818, 1997. 51. Bernardini, M.L. et al. Identification of icsA, a plasmid locus of Shigella flexneri that governs bacterial intra- and intercellular spread through interaction with F-actin. Proc. Natl. Acad. Sci. USA, 86, 3867, 1989. 52. Makino, S. et al. A genetic determinate required for continuous reinfection of adjacent cells on a large plasmid of Shigella flexneri 2a. Cell, 46, 551, 1986. 53. Goldberg, M.B. et al. Unipolar localization and ATPase activity of IcsA, a Shigella flexneri protein involved in intracellular movement. J. Bacteriol., 175, 2189, 1993. 54. Lett, M.-C. et al. virG, a plasmid-coded virulence gene of Shigella flexneri: identification of the virG protein and determination of the complete coding sequence. J. Bacteriol., 171, 353, 1989. 55. Suzuki, T., Lett, M.-C., and Sasakawa, C. Extracellular transport of VirG protein in Shigella. J. Biol. Chem., 270, 30874, 1995. 56. Sandlin, R.C. et al. Avirulence of rough mutants of Shigella flexneri: requirement of O antigen for correct unipolar localization of IcsA in the bacterial outer membrane. Infect. Immun., 63, 229, 1995. 57. Sandlin, R.C., Goldberg, M.B., and Maurelli, A.T. Effect of O side-chain length and composition on the virulence of Shigella flexneri 2a. Mol. Microbiol., 22, 63, 1996. 58. Sandlin, R.C., and Maurelli, A.T. Establishment of unipolar localization of IcsA in Shigella flexneri 2a is not dependent on virulence plasmid determinants. Infect. Immun., 67, 350, 1999. 59. Robbins, J.R. et al. The making of a gradient: IcsA (VirG) polarity in Shigella flexneri. Mol. Microbiol., 41, 861, 2001. 60. d’Hauteville, H. et al. Lack of cleavage of IcsA in Shigella flexneri causes aberrant movement and allows demonstration of a cross-reactive eukaryotic protein. Infect. Immun., 64, 511, 1996. 61. Egile, C. et al. SopA, the outer membrane protease responsible for polar localization of IcsA in Shigella flexneri. Mol. Microbiol., 23, 1063, 1997. 62. Steinhauer, J. et al. The unipolar Shigella surface protein IcsA is targeted directly to the bacterial old pole: IcsP cleavage of IcsA occurs over the entire bacterial surface. Mol. Microbiol., 17, 945, 1999. 63. Monack, D.M., and Theriot, J.A. Actin-based motility is sufficient for bacterial membrane protrusion formation and host cell uptake. Cell. Microbiol., 3, 633, 2001.

Shigella 64. Suzuki T., Saga, S., and Sasakawa, C. Functional analysis of Shigella VirG domains essential for interaction with vinculin and actin-based motility. J. Biol. Chem., 271, 21878, 1996. 65. Miki, H., Miura, K., and Takenawa, T. N-WASP, a novel actindepolymerizing protein, regulates the cortical cytoskeletal rearrangement in a PIP2-dependent manner downstream of tyrosine kinases. EMBO J., 15, 5326, 1996. 66. Shibata, T. et al. Cdc42 facilitates invasion but not the actinbased motility of Shigella. Curr. Biol., 12, 341, 2002. 67. Goldberg, M. B. Actin-based motility of intracellular microbial pathogens. Microbiol. Mol. Biol. Rev., 65, 595, 2001. 68. Loisel, T.P. et al. Reconstitution of actin-based motility of Listeria and Shigella using pure proteins. Nature, 401, 613, 1999. 69. Parsot, C., and Sansonetti, P.J. Invasion and the pathogenesis of Shigella infections. Curr. Topics Microbiol. Immunol., 209, 25, 1996. 70. Sansonetti, P.J., and Mounier, J. Metabolic events mediating early killing of host cells infected by Shigella flexneri. Microb. Pathog., 3, 53, 1987. 71. Donohue-Rolfe, A., Acheson, D.W.K., and Keusch, G.T. Shiga toxin: purification, structure, and function. Rev. Infect. Dis., 13 (Suppl 4), S293, 1991. 72. Brown, J.E., Echeverria, P., and Lindberg, A.A. Digalactosylcontaining glycolipids as cell surface receptors for shiga toxin of Shigella dysenteriae 1 and related cytotoxins of Escherichia coli. Rev. Infect. Dis., 13 (Suppl 4), S298, 1991. 73. Keusch, G.T. et al. Shiga toxin: intestinal cell receptors and pathophysiology of enterotoxic effects. Rev. Infect. Dis., 13 (Suppl 4), S304, 1991. 74. Yavzori, M., Cohen, D., and Orr, N. Prevalence of the genes for Shigella enterotoxins 1 and 2 among clinical isolates of Shigella in Israel. Epidemiol. Infect. 128, 533, 2002. 75. Ryan, K.J. Enterobacteriaceae. In: Sherris Medical Microbiology, An Introduction to Infectious Diseases, 4th Edn. Ryan, K.J., and Ray, C.G., Eds. McGraw-Hill Medical Publishing, NY, 2004. 76. Centers for Disease Control and Prevention (CDC). Disease information: shigellosis. Available at: http://www.cdc.gov/nczved/ dfbmd/disease_listing/shigellosis_gi.html. 2008. Accessed 23 Jul 2008. 77. Kothary, M.H., and Babu, U.S. Infective dose of foodborne pathogens in volunteers: a review. J. Food Safety, 21, 49, 2001. 78. Galanakis, E. et al. Rate of seizures in children with shigellosis. Acta Paediatr. 91, 1001, 2002. 79. Salam, M.A., and Bennish, M.L. Antimicrobial therapy for shigellosis. Rev. Infect. Dis. 13 (Suppl. 4), S332, 1991. 80. Woodward, D.L., and Rodgers, F.G. Surveillance of antimicrobial resistance in salmonella, shigella and Vibrio cholerae in Latin America and the Caribbean: A collaborative project. Can. J. Infect. Dis., 11, 181, 2000. 81. Martin, L.J. et al. Antimicrobial resistance among Salmonella and Shigella isolates in five Canadian provinces (1997 to 2000). Can. J. Infect. Dis. Med. Microbiol., 17, 243, 2006. 82. Sivapalasingham, S. et al. High prevalence of antimicrobial resistance among Shigella isolates in the United States tested by the National Antimicrobial Resistance Monitoring System from 1999 to 2002. Antimicrob. Agents Chemother., 50, 49, 2006. 83. Niyogi, S.K. Increasing antimicrobial resistance–an emerging problem in the treatment of shigellosis. Clin. Microbiol. Infect. Dis., 13, 1141, 2007.

483 84. Lampel, K.A. and Maurelli, A.T. Shigella species. In: Food Microbiology. Fundamentals and Frontiers, 2nd Edn. pp. 216– 227. Doyle, M.P., Beuchat, L.R., and Montville, T.J., Eds. ASM Press, Washington, DC, 2001. 85. June, G.A. et al. Evaluation of the Bacteriological Analytical Manual culture method for the recovery of Shigella sonnei from selected foods. J. AOAC Int., 76, 1240, 1993. 86. Jacobson, A.P. et al. Evaluation of the Bacteriological Analytical Manual (BAM) culture method for the detection of Shigella sonnei in selected types of produce. [poster] FDA Science Forum. February 20–21; Washington, DC, 2002. 87. Warren, B.R. Comparison of conventional culture methods and polymerase chain reaction for the detection of Shigella spp. on tomato surfaces. Masters thesis, University of Florida, Gainesville, FL, 2003. 88. Islam, D. et al. Rapid detection of Shigella dysenteriae and Shigella flexneri in faeces by an immunomagnetic assay with monoclonal antibodies. Eur. J. Clin. Mibrobiol. Infect. Dis., 12, 25, 1993. 89. Islam, D., and Lindberg, A.A. Detection of Shigella dysenteriae type 1 and Shigella flexneri in feces by immunomagnetic isolation and polymerase chain reaction. J. Clin. Microbiol., 30, 2801, 1992. 90. Achi, R. et al. Immunomagnetic separation and PCR detection show Shigellae to be common faecal agents in children from urban marginal communities of Costa Rica. J. Infect., 32, 211, 1996. 91. Achi-Berglund, R., and Lindberg, A.A. Rapid and sensitive detection of Shigella sonnei in feces by the use of an O-antigen-specific monoclonal antibody in a combined immunomagnetic separation-polymerase chain reaction assay. Clin. Microbiol. Infect., 2, 55, 1996. 92. Sansonetti, P.J., Kopecko, D.J., and Formal, S.B. Shigella sonnei plasmids: Evidence that a large plasmid is necessary for virulence. Infect. Immun. 34, 75, 1981. 93. Gamian, A., and Romanowska, E. The core structure of Shigella sonnei lipopolysaccharide and the linkage between O-specific polysaccharide and the core region. Eur. J.  Biochem., 129, 105, 1982. 94. Kimura, A.C. et al. Multistate shigellosis outbreak and commercially prepared food, United States. Emerg. Infect. Dis., 10, 1147, 2004. 95. Andrews, W.H., and Jacobsen, A. Shigella. In: Bacteriological Analytical Manual, 8th Edn. p. 6.01. International AOAC, Gaithersburg, MD, 2001. 96. Wetherington, J. et al. PCR screening and isolation of Shigella sonnei from layered party dip. Lab. Information Bull., 16, LIB no. 4215, 1, 2000. 97. Lindqvist, R. Detection of Shigella spp. in food with a nested PCR method–sensitivity and performance compared with a conventional culture method. J. Appl. Microbiol., 86, 971, 1999. 98. Villalobo, E., and Torres, A. PCR for the detection of Shigella spp. in mayonnaise. Appl. Environ. Microbiol., 64, 1242, 1998. 99. Vantarakis, A. et al. Development of a multiplex PCR detection of Salmonella spp. and Shigella spp. in mussels. Lett. Appl. Microbiol., 31, 105, 2000. 100. Sethabutr, O. et al. Detection of Shigellae and enteroinvasive Escherichia coli by amplification of the invasion plasmid antigen H DNA sequence in patients with dysentery. J. Infect. Dis., 167, 458, 1993. 101. Sethabutr, O. et al. Detection of PCR products of the ipaH gene from Shigella and enteroinvasive Escherichia coli by enzyme linked immunosorbent assay. Diagn. Microbiol. Infect. Dis., 37, 11, 2000.

484 102. Theron, J. et al. A sensitive seminested PCR method for the detection of Shigella in spiked environmental water samples. Water Res., 35, 869, 2001. 103. Lampel, K.A., and Orlandi, P.A. Polymerase chain reaction detection of invasive Shigella and Salmonella enterica in food. Methods Mol. Biol., 179, 235, 2002. 104. Jin, Q. et al. Genome sequence of Shigella flexneri 2a: insights into pathogenicity through comparison with genomes of Escherichia coli K12 and O157. Nucleic Acids Res., 30, 4432, 2002. 105. Warren, B.R., Parish, M.E., and Schneider, K.R. Comparison of conventional culture methods and FTA filtration-nested PCR for the detection of Shigella spp. on tomato surfaces. J. Food Prot., 68, 1606, 2008. 106. Peng, X. et al. Rapid detection of Shigella species in environmental sewage by an immunocapture PCR with universal primers. Appl. Environ. Microbiol., 68, 2580, 2002.

Molecular Detection of Foodborne Pathogens 107. Kakinuma, K., Fukushima, M., and Kawaguchi, R. Detection and identification of Escherichia coli, Shigella, and Salmonella by microarrays using the gyrB gene. Biotech. Bioeng. 83, 721, 2003. 108. Raza, S. et al. Identification and strain differentiation of food and waterborne bacterial theat agents using automated rep-PCR. [poster]. BioDefense Research Meeting, American Society of Microbiology. March 9–12, Baltimore, MD. Available at: http:// www.bacbarcodes.com/posters/BBCI_Poster_2003_Biodefence_ Poster_03_11_03_SR_FINAL.pdf. 2003. Accessed 05 Nov 2004. 109. Lising, M. et al. Identification of Gram-negative bacteria using the DiversiLab system. [poster]. 104th General Meeting, American Society of Microbiology. March 23–27, New Orleans, LA. Available at: http://www.bacbarcodes.com/posters/ ASM_2004_Enteric_Poster.pdf. 2004. Accessed 05 Nov 2004. 110. Food and Drug Administration (FDA). Shigella. In: Bacteriological Analytical Manual, 8th Edn. AOAC International, Arlington, VA, 1998.

35 Vibrio Asim K. Bej

University of Alabama at Birmingham

Contents 35.1 Introduction.................................................................................................................................................................... 485 35.1.1 Classification of Vibrio..................................................................................................................................... 486 35.1.2 Biology and Pathogenesis................................................................................................................................. 486 35.1.2.1 V. cholerae....................................................................................................................................... 487 35.1.2.2 V. parahaemomlyticus..................................................................................................................... 487 35.1.2.3 V. vulnificus...................................................................................................................................... 488 35.1.2.4 Other Vibrios................................................................................................................................... 488 35.1.3 Diagnosis of Vibrio........................................................................................................................................... 488 35.2 Methods.......................................................................................................................................................................... 492 35.2.1 Sample Preparation........................................................................................................................................... 492 35.2.2 Detection Procedures........................................................................................................................................ 493 35.2.2.1 Standard PCR.................................................................................................................................. 493 35.2.2.2 Real-Time PCR................................................................................................................................ 493 35.3 Conclusions and Future Perspectives............................................................................................................................. 495 References.................................................................................................................................................................................. 497

35.1  Introduction Vibrios are typically described as Gram-negative, asporgenic curved, rod-shaped bacteria that are natural inhabitants of fresh, brackish or marine waters. Since the discovery of the first Vibrio sp. (V. choleare) in 1854, more than 45 species have been listed in the recent edition of Bergey’s Manual.1 Most of them are nonpathogenic to humans, but several Vibrio spp. are pathogenic, causing a range of diseases such as gastroenteritis, septicemia, wound infection and ear infection. The severity of infection could be mild to life-­threatening. The primary modes of transmission of these pathogens are potable water, consumption of raw or poorly cooked seafood, or exposure of wounds to warm seawater. However, the primary source of transmission of Vibrio infections is through the consumption of raw or undercooked shellfish. Commonly the disease manifestation following infection by most of the vibrios is gastroenteritis, which is often self-limiting. However, some cases of wound infection and septicemia could also occur particularly to those susceptible individuals who have had liver disease or otherwise immunocompromised often leading to fatal consequences. Although most early studies on seafood-related vibrio infections were from the U.S. and Japan, over the last decade numerous cases have also been described from Asia, Europe, Australia, and New Zealand, suggesting that these pathogens are prevalent in warm coastal waters throughout the world. Although V. cholerae-related cholera disease has not been reported in the U.S. since 1982, frequent incidences and sporadic outbreaks in South America and in

Southeast Asia have been reported. Such outbreaks are often ­widespread during the summer months and occurrences of natural calamities that result in unavailability of clean potable water and poor hygiene conditions. In the U.S., the seafood-related vibrios are prevalent in warm waters of the Gulf of Mexico, although they have also been isolated from Pacific Northwest and Eastern coastal waters. The seafood industry is a major contributor to the local economy of the Gulf of Mexico States and to some extent the Northeast and Pacific Northwest States. Over the last 25 years, the U.S. has experienced several shellfish-related disease outbreaks, with only 4% being due to fecal contaminants (e.g., coliforms, Salmonella sp., Norwalk virus, Hepatitis A), whereas 20% of the diseases and 99% of the deaths were linked to Vibrio spp. present in seafood.2 The microbial pathogens, primarily vibrios are indigenous to the Gulf of Mexico waters and are accumulated by the oysters in their tissues during filterfeeding.3–5 Therefore, consumers inevitably ingest these and other harmful microorganisms when they consume untreated or poorly handled/treated raw oysters. The epidemiological data suggests that the pathogenic vibrios for the incidences and outbreaks of seafood-related diseases in the U.S. and throughout the world are V. parahaemolyticus and V. vulnificus. Many cases of Vibrio-associated gastroenteritis are substantially under-recognized because vibrios are not readily identified in routine stool cultures and may require a timeconsuming lengthy biochemical tests. Therefore the need for rapid and reliable molecular methodologies have been sought and frequently reported. Although the culture-based biochemical and microbiological methodologies are still in 485

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place in many laboratories, slowly but steadily the merits of the molecular approaches are ­realized and gaining popularity in clinical laboratories as well as in the seafood industry. This chapter will elaborate the biology of pathogenic vibrios and molecular approaches for their detection in food.

35.1.1 Classification of Vibrio The family Vibrionaceae under the order Vibrionales includes the members of the genus Vibrio are nonspore forming, Gramnegative bacteria, motile by means of a single polar flagellum, and identified by their unique short rod or curved shape morphology. However two or more polar or lateral flagella have also been demonstrated in Vibrio species, especially when cultured on solid media.1 Also, some members may manifest strait (approximately 1.5–3.0 μm × 0.5 μm) or S-shaped cells when individual cells are joined end-to-end. Because of this morphological diversity, it is necessary to implement physiological and biochemical taxonomic tests for identification and phylogenetic analysis. Vibrio species are facultative anaerobic, manifest both a respiratory (oxygen-utilizing) and a fermentative metabolism. Most of them produce oxidase and catalase, and ferment glucose without producing gas.6 Oxygen has been reported as the universal electron acceptor. Normally, most vibrios inhabit estuarine-, salt- or fresh-waters and can be isolated by culturing on standard growth media such as nutrient agar/broth or media supplemented with required concentrations of NaCl (0.5–3%) and/or a mixture of salts (preferably sea salts). Over the years, the classification of many of the individual vibrio isolates has gone through numerous taxonomic rearrangements. Along with the metabolic characteristics, the DNA G + C composition has played an important role for the taxonomic rearrangement of the Vibrio species. For example, the demarcation between the genus Pseudomonas and Vibrio has been reconsidered based on the physiological and nucleic acids characteristics. The Vibrio species produce acid without forming gas on media supplemented with carbohydrate, therefore considered fermentative, whereas the Pseudomonas species are oxidative. Moreover, the G + C content based on the DNA analysis for Vibrio species is between 39 and 49 mol %; whereas for Pseudomonas species it has been calculated to be 57–67 mol %. A distinct taxonomical separation among the members of the genus Vibrio and Aeromonas, Plesiomonas, Photobaterium, and Lucibacteriu could be somewhat controversial. This is because all of them manifest are fermentative, but do not produce gas (Aeromonas could vary) on media supplemented with carbohydrate; oxidase positive (Photobacterium could vary); and none of them produce diffusible pigment. The primary morphological feature that distinguishes Vibrio species from the rest is that vibrios often manifest round bodies and produce cysts, whereas others do not. Based on these characteristics, Bauman, and Schubert6 have included the following genus, Vibrio, Photobacterium, Aeromonas, and Plesiomonas under the family Vibrionaceae, which was originally described by Veron.7 However, in the most recent edition of Bergey’s Manual, Farmer, and Janda1

Molecular Detection of Foodborne Pathogens

reclassified these genera. In their description, Aeromonas, and Plesoimonas have been classified under different families. This reclassification is based upon the introduction of new methodologies, such as various aspects of proteomics and genomics for the identification of the organisms. In the current Bergey’s Manual, the family Vibrionaceae comprises three genus, Vibrio (Genus I), Photobacterium (Genus II), and Salinivibrio (Genus III).1 Vibrio vulnificus and V. parahemolyticus are natural inhabitants of estuarine and marine waters, respectively, and have been subjected to taxonomic reclassification for many members on this group.

35.1.2  Biology and Pathogenesis After the first description of V. cholerae O1 by Pucini (1854) as the causative agent for cholera in humans, numerous strains have been isolated and taxonomic positions have been assigned. Most vibrios inhabit aquatic environments (freshwater, estuarine, and marine) as free-living organisms or are associated with aquatic animals and plants. However, some vibrios are pathogenic and are isolated from humans when infected and disease symptoms manifested, yet some are pathogenic to aquatic (primarily marine) animals such as fish and eels. Some luminescence vibrios co-exist with marine squid in a symbiotic manner.8 In this chapter, we will focus on vibrios that are primarily pathogenic to humans. Most human pathogenic vibrios reside in freshwater, costal waters (estuarine) and marine environments. The pathogenic vibrios cause infections in humans with the following clinical syndromes (i) gastroenteritis; (ii) wound infection; and (iii) septicemia. The mode of transmission could be through the ingestion of raw or poorly cooked seafood, drinking of potable fresh water causing gastroenteritis or exposure of warm coastal waters to cuts and bruises on the skin leading to septicemia. Some pathogenic vibrios infect normal individuals, while others infect immuncompromised persons particularly those individuals with liver diseases. Most cases of gastroenteritis are self-limiting while the septicemia could be fatal. The Center for Disease Control and Prevention (CDC) in association with the U.S. Food and Drug Administration (FDA) and Gulf Coast States (Alabama, Florida, Mississippi, and Texas) has established Cholera and Other Vibrio Illnesses Surveillance System (COVIS) in 1988, which monitors and reports Vibrio-related illnesses. A comprehensive summary of the human Vibrio isolates since 1998 can be obtained on the CDC website (http://www.cdc.gov/nationalsurveillance/cholera_vibrio_surveillance.html). CDC serotypes all V. parahaemolyticus isolates received from state health departments and screens for cholera toxin production in all V. cholerae isolates. Most states now voluntarily submit reports of clinical cases of Vibrio infections to this site. The primary puropose for the COVIS is to maintain a reliable database of reported cases of Vibrio infections and strain information, and to educate the public about these pathogens, health risks of seafood and other sources of contracting ­disease by these pathogens and how to avoid them.

Vibrio

35.1.2.1  V. cholerae A number of V. cholerae serotypes are potentially pathogenic to humans, whereas nonpathogenic strains have been also been reported. V. cholerae O1 is primarily considered to be causative agent for cholera disease in humans. V. cholerae O1 is subdivided into the most common biotype El Tor or classic; serotype Inaba, Ogawa, or Hikojima; and toxin producing toxigenic or nontoxigenic.9 In 1992, toxigenic V cholerae O139 was first discovered on the Indian subcontinent (Bangladesh) and recognized as another cause of cholera.10 Incidences of V. cholerae O139 have been reported in 1993 in the U.S.11 Due to its similar epidemic potential, monitoring, and surveillance is recommended.11 V. cholerae O1 and O139 are considered to be the pandemic strains. The epidemic strains of V. cholerae are usually the El Tor biotype. Infection with this organism could cause severe watery diarrhea often described as “rice water” stool, vomiting, and muscle cramps, which could lead to severe conditions such as the loss of body weight, loss of normal skin turgor, dry mucous membranes, sunken eyes, lethargy, anuria, weak pulse, altered consciousness, and circulatory collapse and possibly death if not.12 Severe cholera disease causes devastating fluid imbalance leading to dehydration, profound hypokalemia (loss of potassium), metabolic acidosis (from bicarbonate loss), and renal failure. Recently, the V. cholera strains that exhibit resemblance to the clinical strains in their biochemical characteristics but fail to agglutinate in either anti-O1 or anti-O139 sera have been grouped as non-O1/139 V. cholera. These strains of V. cholerae are found in environmental waters, primarily in coastal estuarine waters. These strains could cause choleralike diarrhea that is less severe and are sporadic in nature but rarely been reported to cause outbreaks.13,14 As early as in 1817, V. cholerae seemed to have been causing cholera disease in Asian countries, but spread in the estuarine waters around Africa, Europe, and Americas causing six pandemic outbreaks between 1832 and 1866. The seventh pandemic caused by the V. cholera El Tor biotype was recorded in 1961 spreading from Indonesia to other continents. In 1991, this strain was spread in South America believed to be through ship ballast waters and oceanic current and zooplanktons.15 The outbreak caused by the V. cholera O139 biotype in the early 1990s could be referred to as the eighth cholera pandemic.16,17 The spectrum of illness in cholera includes asymptomatic infection (75%), mild illness (18%), moderate illness (5%), and severe illness (2%).18 35.1.2.2  V. parahaemomlyticus V. parahaemomlyticus is a halophilic estuarine organism that inhabits the coastal waters of virtually all temperate regionsand infections occur primarily during the warmer months of the year. V. parahaemolyticus is the leading cause for gastroenteritis-related illness in humans (irrespective of the health condition). The clinical manifestations of V. parahaemolyticus are acute watery diarrhea and abdominal cramp associated with nausea typically lasting 1–3 days. However severe cases with bloody diarrhea may require hospitalization.

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The primary source of infection by this pathogen is the ­consumption of raw seafood, especially raw oysters. Fujino et al.19 first reported V. parahaemolyticus as a cause for foodborne gastroenteritis in 1950 due to Shirasu food-poisoning in Japan. In 1971, the outbreak of V. parahaemolyticus was first reported in Maryland from the consumption of crabs.20,21 Forty outbreaks of V. parahaemolyticus in the U.S. have been reported to CDC between 1973 and 1998.22,23 These infections occurred primarily during the warmer months and through the consumption of shellfish. In 1997–1998 a relatively large V. parahaemolyticus-related outbreak was reported, affecting over 425 individuals in the U.S. The affected individuals consumed raw oysters harvested from the warm waters of the Gulf of Mexico.22–27 All V. parahaemolyticus strains react with an H antigen. However, several O type and K (capsular) antigen reactive strains have been reported, yet many strains are untypable.28,32 Recently, a newly emerged V. parahaemolyticus O3:K6 serogroup was involved in outbreaks from contaminated oysters harvested from Oyster Bay, NY, and Galveston Bay, TX, in 1998.27 A V. parahaemolyticus-related disease outbreak in Galveston Bay, TX, was the largest reported in the U.S. (416 persons). In 1997, an outbreak caused by V. parahahaemolyticus occurred in the Pacific Northwest, resulting in one death and 209 gatroenteritis-related illnesses. Each of these infected persons had consumed raw oysters.27 Besides gastroenteritis, V. parahaemolyticus can cause wound infection and septicemia. In cases of V. parahaemolyticus-related infections in the U.S. between 1988 and 1997, 59% of patients with gastroenteritis, 34% with wound infections, 5% with primary septicemia, and 2% with other sites of infections from the consumption of raw shellfish were described.22,23 The pathogenic and nonpathogenic strains of V. parahaemolyticus are distinguished primarily based on their ability to produce beta-type hemolysin on Wagatsuma blood agar, which is known as Kanagawa phenomenon.29,30,32–34 This hemolysin is referred to as thermostable direct hemolysin, which is encoded by the tdh gene.29,31 However, there have been cases of illness that were associated with strains that did not produce thermostable direct hemolysin. These strains are referred to as Kanagawa negative, but produce a different hemolysin, the trh-encoded thermostable direct-related hemolysin.22,23 It has been reported that there is a strong association between pathogenicity and the presence of either the tdh gene or the trh gene or both, indicating that in V. parahaemolyticus both of these genes are potential virulence factors.33,34 An increase in the incidence of V. parahaemolyticus around the world was observed in 1996, which was caused by a unique clone of V. parahaemolyticus serotype O3:K6.35,36 It was determined that many of the O3:K6 strains isolated since 1996 contain a filamentous phage, f237, and that this phage contains a unique open reading frame, ORF8. In addition, another group of V. parahaemolyticus strain produces a thermolabile hemolysin, which is encoded by the tlh gene.37 The thermolabile hemolysin does not seem to be associated with pathogenicity, but is present in all V. parahaemolyticus isolates tested so far.38

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Molecular Detection of Foodborne Pathogens

35.1.2.3  V. vulnificus V. vulnificus is halophilic and inhabits coastal warm waters (primarily estuarine water with low ppt salt contents) around the world. In the U.S., V. vulificus is prevalent in the Gulf of Mexico however they are also found in East and West Coast waters. During the warmer months of the year, V. vulnificus counts dramatically increase in coastal waters. In the U.S., V. vulnificus is considered to be the most invasive pathogen among all vibrios causing high fatality among infected individuals, especially those who have had liver disease or are otherwise immunocompromised. The fatality rate could reach 60% in individuals with liver disease and (or) with excess levels of iron in the blood serum.39,40 In addition, open wounds exposed to V. vulnificus in marine waters have been documented as a route of entry for this pathogen, leading to skin breakdown and, in some cases, amputation of the infected body part.40,41 In healthy individuals, infection with V. vulnificus can cause vomiting, diarrhea, and abdominal pains. It was first identified and described by the CDC in 1976.42 Since then it has become the leading cause of seafood-associated deaths in the U.S. The virulence mechanism for V. vulnificus is still not well understood. However, a number of putative virulence factors such as hemolysin, polysaccharide capsule, expression of surface pili, and cellular mechanisms for iron acquisition have been proposed.39,43,44 Among these, it is notable that the capacity for iron acquisition seems to underlie the pathogenicity of many V. vulnificus infections.45A number of studies associate strain virulence with the ability to utilize transferrin-bound iron or with the production of significant amounts of catechol siderophore during infection.46 Moreover, most of the clinical isolates have been identified with thick capsules. Therefore it is proposed that the encapsulated form of V. vulnificus is invasive. Most of the environmental isolates are found to be without capsule and are therefore considered nonpathogenic.47 It has been proposed that V. vulnificus produces a number of enzymes such as lipase, mucinase, hyaluronidase, DNAase, and protease that may facilitate the pathogenesis process.18

and severe abdominal pain.18 This pathogen has been previously referred to as “marine aeromonads,” Group F in England and Group EF6 at CDC.1 Gastroenteritis associated with this pathogen often described as “cholera-like” symptoms. A large outbreak of V. fluvialis related was reported in Bangladesh in 1976–1977.48 The reports of the infection by this pathogen are often from the geographical regions where pathogens causing gastroenteritis are found in feces.1 This pathogen rarely causes septicemia or wound infection.18 The mechanism of pathogenicity has been inflicted upon “enterotoxinlike” molecules tested in rabbit ileal loop.49,50 (iii) V. furnissii is normally found in estuarine environment and has sometimes been isolated from animal feces. Often V. furnissii were isolated from patients along with other pathogenic vibrios. Therefore the role of V. furnisssii as the actual cause for diarrhea remains unclear.1 The pathogenic mechanism by this pathogen is yet to be described. (iv) V. metchnikovii is normally isolated from brackish, marine or fresh water, but rarely from clinical samples. Normally V. metchnikovii is isolated from nonhuman sources such as river, fowl, and other seafood, which could potentially serve as the reservoir for this pathogen to be transmitted to humans.1 It was isolated from the blood of a diabetic individual with acute gall bladder problem.51 V. metschnikovii was isolated from children with diarrhea in Peru and persisted but no subsequent infection was reported.52 The nature of the virulence factor or the mechanism of pathogenesis is not understood. (v) V. hollisae is a halophilic bacteria isolated from sporadic cases of patients with diarrhea or from blood.1 However the pathogenic role has not been firmly established. It could cause septicemia and wound infections.53,54

35.1.2.4 Other Vibrios Besides V. cholerae, V. parahaemolyticus, and V. vulnificus, a few other marine vibrios, although not prevalent, have been reported to cause infections in humans. Some of the halophilic vibrios inhabit brackish waters that have been reported to cause human disease are V. alginolyticus, V. fluvialis, V. furnissii, V. metchnikovii, and V. hollisae. A short description and their possible pathogenic mechanisms are described below:

The conventional culture methods for the detection of vibrios often rely upon the enrichment of the sample and plating on vibrio-specific agar plates followed by a series of other biochemical tests for the confirmation of the results. As a pro-active measure to prevent or reduce vibrio-related incidences and outbreaks, monitoring these pathogens in relevant environmental and food samples for early detection has been recommended. For clinical situations where a patient is suspected to have been infected with a vibrio pathogen, rapid detection, and initiation of treatments regimes are essential. This is particularly important for invasive vibrios such as V. cholerae and V. vulnificus, because these may rapidly progress to fatal consequences.55,56 Many cases of Vibrioassociated gastroenteritis are under-recognized because most clinical laboratories do not routinely use the selective medium, thiosulfate-citrate-bile salts-sucrose (TCBS) agar, for processing of stool specimens unless they are specifically

(i) V. alginolyticus has been isolated from patients with wound infection, although it can cause ear infection too. Howeve,r unlike the other three primary vibrio pathogens, V. alginolyticus rarely causes gastroenteritis.1,18 The virulence mechanism for this pathogen is yet to be described. (ii) V. fluvialis has been known to cause diarrhea, often with bloody diarrhea, fever, vomiting, dehydration,

35.1.3  Diagnosis of Vibrio

Vibrio

instructed to do so. Therefore, a heightened awareness and understanding of appropriate measures, such as implementation of post-harvest treatment and rapid detection methods for seafood products, by the seafood industry, government agencies, clinicians, laboratory technicians, and epidemiologists are essential.57 Vibrio species grow in the presence of relatively high levels of bile salts. Since they are facultative anaerobic and are inhabitants of marine or brackish waters, alkaline conditions along with various concentrations of salts are preferable and essential for their optimum growth. For example, V. parahaemolyticus is strictly halophilic therefore a 2–3% NaCl is necessary for optimum growth. However, for V. vulnificus and other vibrios inhabiting brackish waters, approximately 0.5–1% NaCl is sufficient. Normally, the TCBS agar, which was originally developed for the isolation of pathogenic vibrios is widely used for the isolation of Vibrio species.58 TCBS utilizes alkaline pH (8.6), ox bile, and NaCl (1%) to suppress the growth of nontarget bacteria, including members of the ­family Enterobacteriaceae, the genus Pseudomonas, and Gram-positive bacteria. The stool specimens can be cultured on blood and MacConkey agars, but isolation is enhanced by using TCBS agar.59,60 V. cholerae appears as yellow colonies on TCBS agar. A serological test can be conducted for cholera diagnosis. The serologic conversion, vibriocidal antibody titer of greater than 1:640, suggests a recent infection or a four-fold rise in vibriocidal antibody titer is a relatively reliable indicator. An increase in the titers a couple of weeks after exposure followed by a decrease can often be seen in patients who have had been exposed to V. cholerae.18 V. parahaemolyticus isolates can be maintained on nutrient-rich (Tryptone) agar supplemented with 3% NaCl. However, similar to other vibrios, V. parahaemolyticus may be overlooked if plated on nonselective media; therefore, they are plated on TCBS agar. On TCBS agar V. parahaemolyticus isolates appear as distinct green colonies. Virulent strains of V. parahaemolyticus can be determined by β-hemolysis of red blood cells using Wagatsuma blood agarm,30 although newer methods that use DNA gene probes and PCR have been described.38 The microbiological growth media and conditions for isolation and culture confirmation of V. vulnificus have been reviewed by Harwood et al.4 Normally the enriched cultures of V. vulnificus in oysters or other seafood homogenates can be isolated on TCBS agar. The V. vulnificus appears as green colonies on TCBS agar. Although the TCBS agar is widely used for the isolation of V. vulnificus, V. cholerae and V. parahaemolyticus it has limitations.60 The cellobiose polymyxin B colistin (CPC) and modified-CPC (mCPC) agar media were specifically formulated for the isolation of V. vulnificus in environmental samples.61 These agar media have been shown over 99% efficient in isolating V. vulnificus and are commonly used in most laboratories. Besides the mCPC agar, VVMc agar has shown promising results in the isolation of V. vulnificus.62,63 This agar medium is a modification of VVM agar that does not contain polymyxin B.64

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A comparison of the efficiency of the recovery of V. ­vulnificus from environmental and clinical sources was compared on VVM, VVMc, CC, and TCBS agar. V. vulnificus exhibited highest on VVMc (53%) and lowest on TCBS (42%).62,63 It is important to note that the samples from eels did not support any growth of V. vulnificus on VVMc or CC agar. This suggests that the V. vulnificus Biotype II, which is primarily found in eels, requires different media composition for their growth. The Vibrio species can be identified and characterized using a commercially available API 20E Biolog system. Although the method itself could be useful, this method requires (i) isolation of individual colonies after at least 24 h incubation on TCBS agar; (ii) selection of large number of colonies on TCBS agar for testing, as this agar medium itself is not confirmative. Therefore, this approach is not practical for screening a large number of samples. The application of conventional culture methods followed by a series of confirmative biochemical tests for the evaluation of a large number of food or clinical samples is not convenient.65 Molecular techniques have been gaining popularity for the detection of pathogenic vibrios in coastal waters, seafood, and clinical samples. Besides plate cultures and confirmatory biochemical tests, immunoassays for the detection of V. parahaemolyticus or V. vulnificus are described. However the specificity and the sensitivity of the detection of specific Vibrio species were not reliable. The genetic-based approaches are considerably reliable and often take much less time than the culture-based biochemical testing. DNA-based molecular approaches offer time-efficient specific detection of pathogenic Vibrio species without compromising the required sensitivity. The evolution of the DNA-based molecular approaches occurred in a relatively short period of time and notable achievements in the detection technology have been made. For the molecular approach, a DNA-DNA colony hybridization using either a radiolabeled or a colorimetric tag gene-specific DNA probe was first reported. Yamamoto et al. described the use of a 3.2-kbp segment consisting of the V. vulnificus cytolysin gene (also referred as cthA, hlyA or vvhA, GenBank accession no. M34670) labeled with radionucleotide as a DNA probe, the specificity of which was later confirmed for the detection of environmental and clinical isolates of V. vulnificus.66 Later, an oligonucleotide probe labeled with alkaline phosphatase referred to as VVAP was developed and tested for its specificity for the colorimetric detection of V. vulnificus in environmental samples.64,66 In addition to the detection of V. vulnificus, tlh, and tdh gene specific probes labeled with alkaline phosphatase have been developed for the detection of total and pathogenic stains of V. parahaemolyticus.67–70 The use of the alkaline phosphatase labeled probes in association with Whatman 541 filter paper for colony lifts method of hybridization detection of targeted Vibrio pathogens are relatively inexpensive and takes 2 days to complete. However, this approach is dependent upon the growth of the targeted pathogen on selective agar media. Microbial pathogens in compromised physiological conditions often fail to grow on selective agar media; or

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cluster when high CFU are present in a culture. This leads to either a “false negative” results or undetermined assay. The colorimetric colony hybridization using alkaline-labeled probe is an FDA-recommended approach still practiced primarily by the FDA Seafood Laboratories for the detection of Vibrio pathogens in shellfish. In this context, it is appropriate to mention that the conventional qualitative method of plate culturing and the most probable number (MPN) technique require 4–7 days to complete.65 The recent applications of PCR technology for the detection of Vibrio species have significantly improved the time and specificity necessary for early assessment of the presence of these pathogens in seafood. After a series of reports on the use of conventional PCR technology, real-time PCR became more popular for such purposes and is currently being proposed as a standard detection method for Vibrio species in seafood. For V. cholerae O1 strain, the cholera toxin gene (ctx) has been targeted. However, many strains do not possess the ctx gene, and V. mimicus being closest to V. cholerae also possesses this gene with over 90% similarity. Conventional PCR targeting a 302-bp DNA fragment of the ctx gene for the detection of pathogenic V. cholerae in shellfish has been reported.71 In another study, the V. cholerae classical El Tor strains was detected targeting the hemolysin (hlyA) gene, whereas the rest of the other biotypes were identified targeting a 869-bp segment of the ompU, a 779-bp region of the toxR and a 0.862-bp region of the tcpAI genes.72,73 PCR amplifications by Brasher et al.71 and Rivera et al.72,73 were performed in a multiplexed format in which all targeted genes were amplified together in a single reaction. Multiplexed PCR is a cost effective, rapid detection method of microbial pathogens. Lyon74 described real-time PCR detection of V. cholerae O1, O139, non-O1 and non-O139 strains using oligonucleotide primers and a TaqMan probe targeting a 70-bp segment of the hlyA gene achieving  < 10 CFU level of sensitivity of detection. The detection of toxigenic V. cholerae by TaqMan PCR targeting the ctx gene was validated against the conventional culture method in bottled water, shrimp, milk, and potato salad.75 The detection level of an initial inoculum of 1–2 CFU/g was achieved in 8 h including 6 h of enrichment. The toxigenic V. cholerae in oysters, sediments, and seawater from the Gulf of Mexico was detected with the minimum level of detection  < 10 CFU.76 In this study, the quantitative analysis was determined using an internal amplification control DNA. Since most incidences and outbreaks of V. parahaemolyticus related gastroenteritis have been reported due to the consumption of raw or poorly cooked oysters, the detection of this pathogen has primarily been targeted to seafood. PCR-based detection methodologies have targeted species specific tlh gene, and tdh and trh genes for the detection of pathogenic V. parahaemolyticus. A conventional multiplexed PCR amplification targeting all three genes for the detection of total and pathogenic strains of V. parahaemolyticus in oyster tissue homogenate have been established.37 Later in 1998, the emergence of a new pandemic pathogenic V. parahaemolyticus serogroup O3:K6 prompted the development of

Molecular Detection of Foodborne Pathogens

PCR-based detection of this pathogen. Using conventional PCR methods, Myers et al.77 targeted a 369-bp segment of the ORF8 for the detection of this newly emerged pandemic pathogen. A multiplexed real-time TaqMan PCR detection targeting the species-specific tlh gene and the pandemic strain O3:K6-specific ORF8 DNA segments developed by Rizvi et al.78 has been applied on oysters and waters from the Gulf of Mexico. The detection level was 1 CFU/ml of initial inoculum of V. parahaemolyticus O3:K6 strain in homogenized oyster shell stock or the Gulf of Mexico waters. Other molecular approaches such as the pulse field gel electrophoresis (PFGE) and arbitrarily primed polymerase chain reaction (AP-PCR) of the pandemic V. parahaemolyticus O3:K6 strains and newly emerged O4:K68, O1:K25, O1:K26, and O1:K untypable strains (collectively referred to as the “pandemic group”) targeting the toxRS/new or orf8 DNA segments exhibited a distinct genotypic cluster.79 In their study, the strains belonging to the “pandemic group,” four other O3:K6 strains that did not possess the tdh gene but possessed the toxRS/new sequence. Additionally, three O3:K6 strains that clearly belonged to the pandemic group by PFGE and AP-PCR did not possess the orf8 sequence. Since PCR primers target only a small segment of the targeted DNA/gene, which is subject to sequence polymorphisms, it would be necessary to perform a DNA–DNA hybridization experiment using the entire orf8 DNA segment as a probe and confirm that in fact these three O3:K6 pandemic strains are devoid of this marker gene. A multiplexed PCR was developed by Okura et al.79 targeting a 263-bp segment of the tdh and a 651-bp segment of the toxRS/new sequence that distinguished the pandemic strains from the rest of the V. parahaemolyticus strains. In another study, the multiplexed real-time PCR assay targeting the tlh, tdh, trh, and the orf8 gene segments and four TaqMan probes labeled with four different fluorophores for the detection of species and pathogenic strains of V. parahaemolyticus in oysters and Gulf of Mexico waters was developed.80 The sensitivity of the combined four-locus multiplexed TaqMan PCR was found to be an initial inoculum of 1 CFU V. parahaemolyticus per gram of oyster tissue homogenate following overnight enrichment. This assay was tested on oysters from the Gulf of Mexico exhibiting 17/33 samples that were positive for tlh and 4/33 samples that were positive for tdh. No sample was positive trh or the orf8 DNA segment. In a separate study, TaqMan quantitative PCR (qPCR) was developed targeting the beta subunit of the DNA gyrase (gyrB) gene and tested on 300 oysters collected from the Eastern Sea of China. The sensitivity of detected has been reported to be < 10 CFU per gram of oyster tissue homogenate. Out of the 300 oysters tested, 78 (26%) were positive for V. parahaemolyticus by the conventional culture method and 97 (32.3%) positive by the real-time PCR assay. All culturepositive samples were PCR-positive. However, 19 samples positive by PCR were culture-negative. These results suggest that although real-time TaqMan PCR could be more sensitive than the culture approach by targeting the “injured” or the “physiologically compromised” V. parahaemolyticus cells,

Vibrio

one must be careful as “nonviable” or “dead” cells or “free DNA” from this species could be amplified resulting in a false-positive detection. A multiplexed real-time TaqMan qPCR targeting the tlh, tdh, and trh gene segments with an IAC was recently described for the detection of species-specific and pathogenic strains of V. parahaemolyticus in oysters.81 The study tested 150 strains of V. parahaemolyticus and other Vibrio species for confirmation of the specificity of the selected primers. The tdh was positive for three strains of V. hollisae suggesting that this assay may not be specific when targeted for this gene. The real-time PCR assay was combined with a MPN format to estimate the quantitative values of the targeted pathogens. In some reactions the inhibition of the IAC amplification by the oyster tissue homogenate was noticed. Nonetheless the IAC allows monitoring of V. parahaemolyticus by real-time PCR amplification and provides a quantitative estimation of the targeted pathogen and helps interpret any “false-negative” results due to the inhibition of the reaction by the oyster matrix. For the detection of V. vulnnificus, initially conventional PCR was developed in seeded oyster tissue homogenate by targeting the vvhA gene.82 Then a different region of the same target was used by Brauns et al.83 for the detection of viable but nonculturable cells (VBNC). An alternative target of V. ­vulnificus-specific 23S rDNA segment was used for a nested PCR amplification with the sensitivity of detection of 120 CFU.84 Multiplexed PCR amplification targeting V. vulnificus and other seafoodborne pathogens was also developed.71,72,85 Due to the inhibition of the oyster tissue matrix, often it was necessary to perform two successive rounds of PCR to achieve the required level of sensitivity of detection. Following multiplexed PCR targeting V. ­vulnificus and other shellfish-borne microbial pathogens, a relatively simple confirmatory detection approach utilizing the Covalink™ microtiter-based ­colorimetric detection technology was developed by Lee et al.86 Detection sensitivity of V. vulnificus in seeded oyster homogenates after 3-h enrichment was 100 CFU/g. The conventional PCR led to the development of a rapid and quantitative detection of V. vulnificus in oysters using the real-time PCR using double-stranded DNA binding SYBR Green I fluorescent dye or TaqMan DNA probes.87 The applications of the real-time PCR in various foods have been reviewed by Levin.88 Several recent reviews detailing other real-time applications88 and problems89 are available. Real-time PCR using SYBR Green I dye targeting a 205-bp segment of the vvhA gene was developed by Panicker et al.90 The assay was shown to be specific and an initial inoculum of 1 CFU could be detected after 5 h of enrichment. Furthermore, the relationship between cell number and cycle threshold signal (Ct value) was linear for both enriched and unenriched samples. This indicates that this method can be used for quantitative analysis of V. vulnificus in oysters. Also, the conventional MPN culture method can be combined with the SYBR Green I-based real-time PCR to establish a rapid and cost-effective confirmatory quantitative test. This will also allow detection of viable cells in the samples and avoid

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the possibility of the detection of “nonviable” or “dead” cell in the samples. In addition to the SYBR Green I-based real-time PCR assay, TaqMan probe was also used for the detection of V. vulnificus in oysters targeting a 100-bp DNA fragment of the vvhA gene.91 The sensitivity of detection was determined to be 100 CFU/g of oyster tissue homogenate. This study showed that a much higher cell counts (107 CFU/g) are necessary for the detection by the TaqMan assay. Three different primer sets and their internal TaqMan probes targeting the vvhA gene have been compared by realtime PCR method.92 Although all three primer sets exhibited positive amplification of all 81 V. vulnificus strains tested, a few other Vibrio species exhibited positive Ct values indicating false-positive detection by one of the three sets of primer. This is most likely due to the smaller amplicon size leading to the formation of PCR artifacts rather than the specificity of the targeted gene for V. vulnificus. Identification of clinical strains of V. vulnificus was developed using real-time PCR and SYBR Green I targeting the vulnibactin (viuB) gene, the product of which is involved in chelating iron from the outer environment and transporting it into the cytoplasm.90 In this study, a 504-bp segment of the viuB gene and a 205-bp segment of the vvhA gene were used in a multiplexed format for the detection of total and clinical strains of V. vulnificus in oysters with a sensitivity level of an initial inoculum of 1 CFU/g following overnight enrichment. Application of this assay on oysters collected from the Gulf of Mexico exhibited 51% of the samples positive for vvhA and 15% of them were positive for viuB. During the winter months, V. vulnificus has been reported to remain in a VBNC state.93,94 Therefore the conventional culture techniques may detect this pathogen when present in this state. However, the genetic-based technologies targeting species-specific genes have the potential to detect V. vulnificus and other Vibrio species during their existence in the VBNC state. Campbell and Wright91 used the TaqMan PCR probe targeting vvhA gene on a VBNC culture and were able to achieve positive detection at 107 CFU V. vulnificus per gram of oyster tissue homogenate. Detection of this high level of CFU coincides with the previous observation by Brauns et al.83 Similarly, Ward, and Bej80 showed relatively high Ct values on cold-stressed VBNC V. parahaemolyticus cultures with low CFU per ml as compared to the cultures maintained in normal temperature indicating that high cell counts are necessary for the detection of VBNC cultures by PCR methods. Detection of viable cells could be better determined by targeting the mRNA of metabolically active signature genes of a pathogen. Studies for such detection have been documented for several Vibrio species. The application of reverse transcriptase PCR (RT-PCR) on vvhA transcript in VBNC V. vulnificus showed sustained expression during this physiological state.95 In a separate study, RT-PCR on the pathogenic genes in VBNC V. parahaemolyticus cultures were shown to remain transcriptionally silent.96,97 Therefore it may be wise to select an essential house-keeping gene for PCR amplification

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detection of VBNC cultures. Sun et al.98 reported that VBNC cultures of V. harveyi do not cause infection to the host (fish), but when resuscitated the cultures become infectious. This result suggests that although the virulence potential for the pathogenic Vibrio species is diminished during the VBNC state, they remain pathogenic when resuscitated. This observation is supported by a previous study, in which VBNC cultures of V. alginolyticus and V. parahaemolyticus were noninfectious to mice, but become infectious when resuscitated.99 An RT-qPCR analysis of the VBNC V. cholerae culture showed expression of an essential protein synthesis and a stress responsive gene, relA, and rpoS, respectively.100 This study indicates that careful selection of target genes for the detection of the viable Vibrio cells in environmental and food samples are essential. Although the RT-PCR approach seems to be an effective method for the detection of VBNC Vibrio species, challenges related to the inhibition of the reaction by the sample matrix, effectiveness of the reverse transcriptase reaction, and possibility of a positive detection signal from the nonviable cells in the sample remain. In addition the issues relevant to the bacterial viability, the “true” existence of VBNC cells in nature and in seafood remain controversial.

35.2  Methods 35.2.1 Sample Preparation Sample preparation remains the crucial step for successful detection of pathogenic Vibrio species in seafood samples by PCR amplification. The primary foci are to avoid the inhibition of PCR leading to a “false negative” detection; achieve sensitivity of detection that complies with the Interstate Shellfish Sanitation Conference (ISSC) guidelines, and avoid detection of nonviable/dead cells, which could lead to a “false-positive” detection. The current general consensus of all procedures is to include a sample enrichment step prior to PCR amplification, which helps to achieve the aforementioned objectives. An enrichment growth medium suitably selective for V. cholerae has not been documented. Currently Alkaline Peptone Water (APW)52 is used for enrichment, in which V. cholerae can grow to a sufficient density for PCR amplification detection within 6–8 h. Since APW is not selective, it supports other competing microorganisms to grow simultaneously affecting the growth rate for V. cholerae. However, Depaola, and Hwang68 have shown that overnight enrichment resulted into an effective isolation of V. choleae. Longer enrichment may be necessary for better recovery of V. cholerae and other pathogenic vibrios. Injured cells are the result of samples subjected to post-harvest treatments such as freezing and thawing or high-pressure treatment. In addition, the incubation of raw oyster shellstock in APW at 42°C for 6–8 h has been shown to be effective for the isolation of V. cholerae and is currently recommended.69 The general enrichment protocol and conventional culture-based identification and enumeration methods for

Molecular Detection of Foodborne Pathogens

pathogenic Vibrio species currently recommended by the FDA Bacteriological Analytical Manual101 are as follows: (i) V. cholerae: (1) Dilute raw oyster shellstock to 1:100 ratio of oyster to APW. Transfer 25 g of sample into a tared jar (preferable capacity approximately 500 ml). Products such as seafood or vegetables may be blended or cut into small pieces with sterile scissors or knife. (2) Add 225 ml APW (pH 8.4) to the jar. Thoroughly mix the sample or blend 2 min at high speed. (3) Incubate APW at 35 ± 2°C for 6–8 h. Re-incubate the jar overnight if desired. (4) For analysis of raw oysters, it is recommended that a second tared jar with 25 g of product plus 2,475 ml APW be incubated 18–21 h at 42 ± 0.2°C in a water bath.65,68 Comment: An enumeration technique by MPN may also be performed if desired. A sample from the MPN positive tubes can be subjected to PCR amplification targeting the species-specific gene(s) for the detection and establishment of a quantitative assessment of the targeted pathogen (see Section 35.2.2). (5) For conventional culture-based detection, the cultures can be ten-fold serially diluted in APW and applied onto TCBS agar or mCPC agar plates by spread plating or streaking the pellicle of the sample with a 1-mm sterile loop in a manner that will yield single colony isolation. TCBS agar is incubated overnight (18–24 h) at 35 ± 2°C. Incubate mCPC and CC overnight at 39–40°C or at 35–37°C. V. cholerae on TCBS agar can be identified by the morphology and texture such as large (2–3 mm), smooth, yellow, and slightly flattened with opaque centers and translucent peripheries. Typical colonies of V. cholerae on mCPC or CC agar are small, smooth, opaque, and green to purple in color, with a purple background on extended incubation. (6) For biochemical identification, colonies from crowded plates must be streaked on nonselective agar (T1N1, T1N3, or TSA-2% NaCl agar) for purity. Incubate overnight at 35 ± 2°C and proceed with identification using a single isolated colony. (7) Subculture three or more typical colonies from each plating medium to T1N1 agar slants or motility test medium stabs. Incubate slants or stabs overnight at 35 ± 2°C. (ii) V. vulnificus: (1) For fish samples the surface tissues, gills, and for shellfish samples the shellstock (meat and liquor) from 12 to 15 animals of approximately 50 g of weight are analyzed. The samples are pooled and blended at high speed for 90–120 sec

493

Vibrio

(if necessary with pulses) until the ­homogenate is formed. For crustaceans such as shrimp or crayfish the entire animal is used. However if the animal is too large then the central portion, including gill and gut are used. (This step is the same for V. parahaemolyticus). (2) Tissue homogenates (preferably three tubes each of 1-ml) are inoculated in 100 ml APW and incubated at 35–37°C for 18–24 h in a rotary shaker set at 150–170 rpm. (3) For plate culture analysis, a 3-mm loopful from the top 1 cm of APW tubes is streaked onto mCPC or CC selective agars. Incubate mCPC and CC agars overnight at 39–40°C (alternatively 35–37°C if 39–40°C not available). On either agar, colonies appear as round, flat, opaque, yellow, and are 1–2 mm in diameter. (4) Along with the plate cultures, a three-tube MPN method as described above for the V. cholerae can be conducted. The MPN-positive tubes can be subjected to PCR amplification for the confirmative and quantitative detection of V. vulnificus. (5) Individual colonies can be picked by using a sterile toothpick and resuspended in 10 μl sterile water and boiled for 10 min to release DNA. The released DNA can be subjected to PCR amplication using V. vulnificus gene-specific primers (vvhA) for detection. (iii) V. parahaemolyticus: (1) The enrichment step is as for V. vulnificus described above. After enrichment, the culture is streaked on TCBS agar using the method described above and grown at 35–37°C overnight. V. parahaemolyticus colonies appear as round, opaque, green or bluish colonies, 2–3 mm in diameter on TCBS agar. V. parahaemolyticus colonies can be differentiated from V. alginolyticus colonies, which are large, opaque, and yellow. Most strains of V. parahaemolyticus will not grow on mCPC or CC agar. If growth occurs, colonies will be green-purple in color due to lack of cellobiose fermentation. (2) An aliquot of the culture can be subjected to three-tube MPN enrichment, if desired. The method is described above. (3) Biochemically, V. parahaemolyticus and V. vulnificus are phenotypically similar, but can be differentiated by differences of the ONPG, salt-tolerance, cellobiose, and lactose reactions. (4) The urease positive for V. parahaemolyticus indicates the potentially pathogenic strain. (5) Either the enriched or the MPN enriched cultures could be subjected to PCR amplification detection of species and pathogenic strains of V. parahaemolyticus.

35.2.2  Detection Procedures 35.2.2.1 Standard PCR Principles: Standard PCR assays based on agarose gel format have been developed for identification of pathogenic V. cholerae,71,74 V. vulinificus71 and V. parahaemolyticus37 (Table 35.1). We present below step-wise PCR protocols for Vibrio detection with primers from these authors. Procedure: (1) Prepare PCR master mix containing 200 µM of each dNTP, 1 µM of each primer (see Table 35.1), 2.5 U of Taq DNA polymerase, 1 × PCR buffer [50 mM Tris·Cl pH 8.9, 50 mM KCl, 2.5 mM MgCl2 (referred to as buffer C)], and sterile distilled water to 25 or 50 μl per reaction (depending upon the make and model of the thermal cycler). Transfer an appropriate aliquot into a PCR tube. Add 1–3 µl of boiled DNA sample per reaction. Include a tube with purified genomic DNA as positive control and a tube with master mix (plus distilled water) as negative control. (2) Perform PCR amplification in a thermocycler with one cycle of 94°C for 3 min; 25–30 cycles of 94°C for 0.5–1 min, 55–60°C for 0.5–1 min, and 72°C for 0.5–1 min; and one cycle of 72°C for 5–7 min followed by the soak cycle at 4–10°C until gel electrophoresis analysis. (3) Separate PCR product (5 μl) on a 1–1.2% (w/v) agarose gel in 1 × TAE (pH 8.2) buffer containing ethidium bromide (0.5 μg/ml) at 5 V/cm. Visualize the specific products under UV light. Note: If further confirmation is necessary Southern blot DNA–DNA hybridization can be performed using the standard method described by Ausubel et al.102 The probes specific for the hybridization detection of the amplified DNA are described in Table 35.1. 35.2.2.2 Real-Time PCR Principles: Several real-time PCR assays based on TaqMan have been reported, which facilitate improved detection and identification of pathogenic V. cholerae,74,76 V. vulinificus,92 and V. parahaemolyticus77–80 (Table 35.1). (i) V. cholerae [Non-classical hlyA (El Tor, O139; non-El Tor, non-O139)]: (1) Prepare PCR mix (50 μl) consisting of 1 μl TaqMan buffer A (Perkin Elmer Applied Biosystems, carlsbad, CA); 5 mM MgCl2; 200 μM (each) of dATP, dCTP, and dGTP; 400 μM dUTP; 0.02 μM TaqMan probe specific for the nonclassical hlyA gene (Table 35.1); 0.3 μM (each) V. cholerae specific primers targeting the nonclassical hlyA gene (Table 35.1); 1 U of AmpErase uracil N-glycosylase; 2.5 U of AmpliTaq Gold DNA polymerase; and DNA sample (2.5 μl, 100 ng/μl)

494

Molecular Detection of Foodborne Pathogens

Table 35.1 Description of Primers and Probes for PCR Amplification Detection of Pathogenic Vibrio Species Microorganism

Targeted Gene

Primers/Probes

Tm (°C)

Amplicon Size (bp)

Reference

Conventinal PCR ctxA (Classical El Tor)

L-CTX: 5′-ctcagacgggatttgttaggcacg-3′ R-CTX: 5′-tctatctctgtagccggtattacg-3′ P-CTX2: 5′-attccatactccccaaatata-3′

74 70

302

71

Non-classical hlyA (El Tor, O139; non- El Tor, non-O139)

Forward 5′-tgcgttaaacacgaacgcat-3′ Reverse 5′-aagtcttacattgtgcttgggtca-3′ Probe: 5′-tcaaccgatgcgattgcccaaga-3′

58 59

70

74

V. vulnificus

vvhA (species specific)

L-CTH: 5′-ttccaacttcaaaccgaactatgac-3′ R-CTH: 5′-gctactttctagcattttctctgc-3′ P-CTH: 5′-gaagcgcccgtgtctgaaactggcgtaacg-3′

70 68

205

71

V. parahaemolyticus

tlh (species specific)

L-TL:5′-aaagcggattatgcagaagcactg-3′ R-TL:5′-gctactttctagcattttctctgc-3′ P-TL: 5′-acggacgcaggtgcgaagaacttcatgttg-3′

70 68

450

38

tdh (pathogenic strain)

L-TDH:5′-gtaaaggtctctgacttttggac-3′ R-TDH:5′-tggaatagaaccttcatcttcacc-3′ P-TDH: 5′-gcggtgtctggctstssgsgcggtcssttct-3′

68 68

269

trh (pathogenic strain)

L-TRH:5′-ttggcttcgatattttcagtatct-3′ R-TRH:5′-B-cataacaaacatatgcccatttccg-3′ P-TRH: 5′-aaaactgaatcaccagttaacgcaatcgtt-3′

66 60

500

orf8 (pandemic O3:K6 serotype)

F-O3MM824 5′aggacgcagttacgcttgatg-3′ R-O3MM1192 5′-ctaacgcattgtccctttgtag-3′

64 64

369

V. cholerae

Real-Time PCR ctxA (Classical El Tor)

Forward: 5′-tttgttaggcacgatgatggat-3′ Reverse: 5′-accagacaatatagtttgacccactaag-3′ Probe: FAM-5′-tgtttccacctcaattagtttgagaagtgccc-3′ BHQ

Non-classical hlyA ( El Tor, O139; non- El Tor, non-O139)

Forward 5′-tgcgttaaacacgaacgcat-3′ Reverse 5′-aagtcttacattgtgcttgggtca-3′ TaqMan probe FAM 5′-tcaaccgatgcgattgcccaaga-3′ TAMRA

V. vulnificus

vvhA

F-vvh785 ttccaacttcaaaccgaactatgac-3′ R-vvh990 attccagtcgatgcgaatacgttg-3′ P-vvh874 FAM-5′-aactatcgtgcacgctttggtaccgt-3′ BHQ-1

V. parahaemolyticus

tlh (species specific)

L-TL:5′-aaagcggattatgcagaagcactg-3′ R-TLH:5′-gctactttctagcattttctctgc-3′ P-TL: 5′-TEXR-aagaacttcatgttgatgacact-BHQ2-3′

tdh (Pathogenic strain)

L-TDH:5′-gtaaaggtctctgacttttggac-3′ R-TDH:5′-tggaatagaaccttcatcttcacc-3′ P-TDH:5′-CY5-attttacgaacacagcagaatga-3′ Iowa Black-RQ-3

68 68

269

trh (pathogenic strain)

L-TRH:5′-ttggcttcgatattttcagtatct-3′ R-TRH:5′-B-cataacaaacatatgcccatttccg-3′ P-TRH:5′-TET-tatttgtygttagaaatacaacaat-BHQ1-3′

66 60

500

orf8 (pandemic O3:K6 serotype)

F-O3MM824 5′-aggacgcagttacgcttgatg-3′ R-O3MM1192 5′-ctaacgcattgtccctttgtag-3′ P-ORH8: 5′-6-FAM-aagccattaacagttgaaggcgttgact-BHQ1-3′

64 64

369

V. cholerae

84

76

58 59

70

74

66 70

205

92

450

78,80

77

Y = C or T; M=A or C; R = G or A; F = forward primer; R = reverse primer; P = probe. TexR, sulforhodamine 101 (Texas Red) fluorescent dye; BHQ2, Black Hole-2 quencher dye; 6-FAM, 6-fluorescein fluorescent dye; BHQ1, Black Hole-1 quencher dye; Cy5, carbocyanine fluorescent dye; Iowa Black-RQ, Iowa Black quencher dye; TET, tetrachloro-6-carboxyfluorescein fluorescent dye; ROX, 6-carboxy-X-rhodamine; BHQ-2, Black Hole-2 quencher dye; FAM, 6-fluorescein; BHQ-1, Black Hole-1 quencher dye. Tm (°C) = 2(A + T) + 4(G + C). bp = base pair.

495

Vibrio

in 0.2 ml of MicroAmp Optical reaction tubes with MicroAmp Optical tube caps (PE Applied Biosystems). (2) Keep the PCR mixture at 50°C for 5 min and then denature at 95°C for 10 min. Carry out 40 cycles of 95°C for 20 sec followed by 60°C for 1 min using the ABI Prism 7700 sequence detection system (PE Applied Biosystems) using an ABI Prism 7700 sequence detection system and SOS (version 1.7) application software (PE Applied Biosystems). Other real-time PCR thermocyclers may also be used. (ii) V. cholerae (Classical El Tor): (1) Prepare PCR mix (25 μl) consisting of 1 × PCR Buffer (Invitrogen, Carlsbad, CA), 5 mM MgCl2, 0.2 mM each of the dNTPs, 1.25 U of Platinum Taq polymerase (Invitrogen), 250 nM primers targeting the nonclassical hlyA gene (Table 35.1), 100 nM TaqMan probe specific for the ctx gene (Table 35.1) and DNA sample (1.5 μl, 100 ng/μl). (2) Perform real-time PCR with one cycle of 94°C for 2 min; 45 cycles of 94°C for 10 sec, 63°C for 30 sec (with the instrument optics on) using a Cepehid Smart Cycler™ II with all default analysis parameters except that the manual threshold fluorescence units setting is adjusted to 10. Other real-time PCR thermocyclers may also be used. (iii) V. vulnificus: (1) Prepare PCR mix (25 μl) consisting of 2.5 μl of 10 × PCR buffer (200 mM Tris-Cl [pH 8.4], 500 mM KCl), 5 μl of 25 mM MgCl2, 200 μM ­concentrations of dNTPs, 0.24 μM TaqMan probe specific for the vvhA gene, 0.4 μM of the primers targeting the vvhA gene, 1.5 U of Taq DNA polymerase, and DNA sample (1.5 μl, 100 ng/μl). (2) Perform real-time PCR with one cycle of 94°C for 2 min; 40 cycles of 94°C for 15 sec, 58°C for 15 sec, and 72°C for 20 sec using a Cepehid Smart Cycler™ II with all default analysis parameters and the set cycle threshold set at 30. (iv) V. parahaemolyticus: For amplification of V. parahaemolyticus species and pathogenic strains a multiplexed PCR is recommended with four sets of primers targeting the tlh, tdh, trh, and orf8 genes and four gene specific TaqMan probes (Table 35.1). (1) Prepare PCR mix (25 μl) consisting of 1 × PCR buffer (20 mM Tris-Cl [pH 8.4], 50 mM KCl), 5 mM MgCl2, 400 μM of each dNTPs, 0.4 μM of each of the eight gene-specific oligonucleotide primers, 0.2 μM of each of the four fluorescencelabeled TaqMan probes, 100 ng bovine serum

albumin, 2 U of Platinum Taq DNA polymerase (Invitrogen) (2) Perform real-time PCR with one cycle of 94°C for 2 min; 45 cycles of 94°C for 20 sec, 56°C for 20 sec, and 72°C for 30 sec using a Cepehid Smart Cycler™ II. The increase in fluorescence for each channel is measured and recorded during the annealing step of each cycle. The cycle threshold (Ct) value is set at 15 fluorescence units. Note: Recently Nordstrom et al.81 described the use of an IAC DNA, which is co-amplified with the targeted gene(s). The IAC avoids the “false-negative” detection and helps determine the quantitative estimation of the initial copies of the targeted gene(s). Since the seafood or the environmental water samples for the detection of Vibrio species are enriched, the quantitative estimation of the targeted pathogens can not be determined accurately. Therefore the use of the IAC for the detection of Vibrio species is restricted to confirm the integrity of the PCR amplification process.

35.3 Conclusions and Future Perspectives Historically V. cholerae within the Vibrioneaceae family received the most attention for cholera outbreaks including several pandemic outbreaks and it continues be a problem primarily in South America, Southeast Asia, and Africa. However, since the last report of the identification of pathogenic strains in the Gulf of Mexico water, this pathogen has not been a problem in the U.S. It is important to note that although the O1 serogroup has not been a problem in the U.S., non-O1 V. cholerae-related diarrhea has been reported. Therefore it is necessary to continuously monitor V. cholerae in food and waters by using rapid detection technologies such as real-time PCR or DNA-microarray to ensure the safety of public health. In contrast, V. parahaemolyticus and V. vulnificus have been a constant problem in the U.S. as well as other countries such as Japan, Southeast Asia, South America and some countries in Europe. Therefore efforts have been devoted to explore rapid, reliable, sensitive, and cost-effective detection technologies by using molecular approaches. During the last two decades, the progress in such detection techniques has been phenomenal. The molecular technologies for the detection of pathogenic vibrios have started with the colorimetric colony blot hybridizations using species-specific DNA probes. The methods were relatively successful in meeting the detection compliances and have been adopted by the FDA seafood laboratories. The second generation molecular techniques have to take advantage of PCR amplification technology. During the first phase of this effort, conventional PCR methodologies were successfully explored for identification of species-specific targeted genes and optimization of the PCR parameters. The conventional PCR approach led to the second phase of PCR-based

496

detection of the pathogenic vibrios, which is the use of the real-time PCR either using the TaqMan probes or a doublestranded DNA-binding fluorescent dye such as SYBR Green I or the Eva Green. The TaqMan PCR method is much faster than conventional PCR and the amplification of the targeted genes can be often monitored in real-time as each cycle of amplification progresses. However, the SYBR Green I-based PCR approach although rapid when compared with the conventional PCR, requires completion of sufficient numbers of amplification cycles at which point a DNA melt-curve analysis is performed to confirm amplification of the targeted gene segments. In that sense, unlike the TaqMan PCR, the SYBR Green I-based PCR method of detection can not be referred as real-time PCR. The transition from the conventional PCR to either TaqMan real-time PCR or the SYBR Green I-based PCR required some optimization of the reaction parameters such the length of the amplicons, reagents concentrations and the shorter PCR temperature cycling parameters. This has led to many studies redesigning the primer locations on the targeted genes, which had been previously established by conventional PCR methods. Besides the PCR approach, a number of reports have taken the advantage of other molecular approaches such as DNA microarray,90 loop-mediated amplification or the helicase dependent isothermal amplification (Mojib and Bej, unpublished data). The DNA microarray offers detection of numerous microbial pathogens simultaneously on a single glass slide with immobilized gene-arrays. However, the process is based on the principles of DNA–DNA hybridization requiring multiple steps of wash cycles followed by computer analysis of the positive hybridization results. Therefore this approach does not offer a real advantage at this time until an automated method is developed. The isothermal amplification approach seems to be the next generation detection technology of the Vibrio species. A few reports have already shown the promise that the isothermal technology offers a cost-effective method of detection without the use of a relatively expensive real-time PCR thermocycler.103,104 However the current format of the loop-mediated isothermal amplification technology requires rather a complex method of primer selection. The other approach of the use of the thermostable helicase-dependent isothermal amplification technology (HDA assay) offers a relatively simple approach of primer selection and isothermal amplification approach. The HDA can be purchased in the kit format from BioHelix, Inc. (www.biohelix.com). One of the challenging issues that yet to be solved is a “realistic” quantitative analysis of Vibrio species and the pathogenic strains in shellfish and other seafood without an enrichment step. This is a rather complex because of the facts that (i) ISSC guidelines recommend the minimum level of specific Vibrio species as a standard for “safe/consumable” shellfish. This also applies to post-harvest treated oysters; (ii) due to the complex tissue matrix, the detection of the ISSC-recommended minimum level of Vibrio pathogens without enrichment is often difficult to achieve; (iii) without enrichment there is a possibility of the detection of dead

Molecular Detection of Foodborne Pathogens

cells or cell-free DNA from the targeted pathogens; (iv) an ­enrichment step may not accurately reflect the initial number of cells in the sample. However, enrichment ensures the detection of viable cells in the samples. To address these issues, a RT-PCR was conducted in which successful amplification of vvhA mRNA allowed the investigators to confirm the detection of viable pathogens.95 However, this approach can be better achieved when applied to the enriched samples and be technically challenging due to relatively short half-life of most prokaryotic mRNA. A quantitative assessment of the pathogenic Vibrio species can be better addressed by combining the conventional MPN method with the real-time PCR approach.105,106 The conventional MPN is specifically applicable to the PHP (Post Harrest Processed) oysters, which often requires process validation, which is often labor-intensive and expensive. The detection of targeted Vibrio species in the MPN-positive tubes by real-time PCR is rapid, reliable, and saves valuable time for subsequent lengthy timeconsuming analysis for confirmation. However, the MPN/ real-time PCR approach requires handling numerous culture tubes and many real-time PCR and could be somewhat expensive. The other issue with Vibrio species is that during the winter months, they seem to enter into VBNC state, which often can not be detected by standard microbiological culture methods.93 Since the direct detection of low levels of Vibrio species in seafood, especially in shellfish tissue without enrichment is often not-reliable leading to “false-negative” detection, revival followed by enrichment of the VBNC cells may be necessary. It is a relatively difficult issue and more attention is needed to come up with a reliable approach to achieve detection of this pathogen when present in the VBNC state. The next generation of nucleic acid detection of targeted genes from Vibrio species could be the application of biosensor technology including nanotechnology.107 This approach is based on the principles of DNA–DNA hybridization on a monolayer of single-stranded DNA probes, which are layered on silicon microcantilever. The positive hybridization of the probe DNA with the targeted gene amplicons can be detected label-free by measuring the changes of the tension in the monolayer as a result of hydration. The tension is changed rapidly when the single-stranded probes on the monolayer interact with the complementary single-stranded amplicons forming hybrids by the nucleic acid base-pair rules. This approach is very sensitive and detection at the femtomole quantity of DNA has been reported. This approach is currently suggested for the detection of multiple gene targets in a microarray format. Molecular detection of Vibrio species using real-time PCR methods is rapid, specific, and could be cost-effective. Also, the PCR-based methods enable us to detect total and pathogenic (clinical) strains of Vibrio species simultaneously in a single reaction. This has been one of the major advantages over conventional culture techniques where a series of biochemical tests are necessary to confirm the species as well as the pathogenic strain. While the quality of the reagents

Vibrio

and the instrumentation for PCR amplification methods have been significantly improved, simultaneously the reliability of the detection of Vibrio species in seafood and clinical samples have become more achievable. However, three aspects of detection (i) a sample processing method that will yield sufficiently pure targeted genomic DNA for inhibition-free amplification of the targeted genes; (ii) a “true” sensitivity of detection without the sample enrichment; and (iii) detection of viable “live” cells in shellfish and other seafood samples need further improvements. In addition, a consensus primer/ probe combination for the detection of Vibrio species and the pathogenic strains should be evaluated and established by an independent research entity so that the application of a “gold standard” approach can be offered to the seafood industry and clinical laboratories for a rapid and effective detection of Vibrio pathogens. This will help the seafood industry in the routine monitoring of this pathogen in seafood, especially in PHP oysters and other shellfish thereby ensuring seafood safety, consumer confidence and protection of public health. Also a consensus detection method will enable clinicians to detect this pathogen in patients with infection rapidly and reliably.

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497 12. Pierce, N.F. and Mondal, A. Clinical features of cholera. In: Cholera, p. 209. Barua, D. and Burrows, W. (eds.). WB Saunders Company, PA, 1974. 13. Morris, J.G. Jr. Vibrio cholerae O139 Bengal: emergence of a new epidemic strain of cholera. Infect. Agents. Dis., 4, 41, 1995. 14. Sharma, C. et al. Molecular analysis of non-O1, non-O139 Vibrio cholerae associated with an unusual upsurge in the incidence of cholera-like disease in Calcutta, India. J. Clin. Microbiol., 36, 756, 1998. 15. Faruque, S.M. et al. Epidemiology, genetics, and ecology of toxigenic Vibrio cholerae. Microbiol. Mol. Biol. Rev., 62, 1301, 1988. 16. Berche, P. et al. The novel epidemic strain O139 is closely related to the pandemic strain O1 of Vibrio cholerae. J. Infect. Dis., 170, 701, 1994. 17. Swerdlow, D.L. and A.A. Ries. Vibrio cholerae non-O1—the eighth pandemic? Lancet 342(8868), 382, 1993. 18. Daniels, N.A. and Shafaie, A. A review of pathogenic Vibrio infections for clinicians. Infect. Med., 17, 665, 2000. 19. Fujino, T. et al. On the bacteriological examination of shirasufood poisoning. Med. J. Osaka Univ., 4, 299, 1953. 20. Dadisman, T.A. Jr. et al., Vibrio parahaemolyticus gastroenteritis in Maryland. I. Clinical and epidemiologic aspects. Am. J. Epidemiol., 96, 414, 1972. 21. Dadisman, T.A. et al. Vibrio parahaemolyticus gastroenteritis in Maryland—clinical and epidemiologic aspects. Am J Epidemiol., 96, 414, 1973. 22. Daniels, N.A. et al. Vibrio parahaemolyticus infections in the United States, 1973–1998. J. Infect. Dis., 181, 1661, 2000a. 23. Daniels, N.A. et al. Emergence of a new Vibrio parahaemolyticus serotype in raw oysters: prevention quandary. JAMA, 284, 1541, 2000b. 24. CDC. Gastroenteritis caused by Vibrio parahaemolyticus aboard a cruise ship. MMWR. 27, 67, 1978. 25. CDC. Foodborne diseases active surveillance network, 1996. MMWR, 46, 258, 1997. 26. CDC. Outbreak of Vibrio parahaemolyticus infections associated with eating raw oysters—Pacific Northwest, 1997. MMWR, 47, 457, 1998. 27. CDC. Outbreak of Vibrio parahaemolyticus infection associated with eating raw oysters or clams harvested from Long Island Sound—Connecticut, New Jersey, and New York, 1998. MMWR, 48, 48, 1999. 28. Miwatani, T. et al. Antibody titers against the thermostable direct hemolysin in the sera of patients with Vibrio parahaemolyticus infection. Kansenshogaku Zasshi 50, 46, 1976. 29. Miyamoto, Y. et al. In vitro hemolytic characteristic of Vibrio parahaemolyticus: its close correlation with human pathogenicity. J. Bacteriol., 100, 1147, 1969. 30. Wagatsuma, S. A medium for the test of the hemolytic activity of Vibrio parahaemolyticus. Media Circle, 13, 159, 1968. 31. Honda, T. and Iida, T. The pathogenicity of Vibrio parahaemolyticus and the role of the thermostable direct haemolysin and related haemolysins. Rev. Med. Microbiol., 4, 106, 1993. 32. Honda, T. et al. Evidence of immunologic cross-reactivity between hemolysins of Vibrio hollisae and Vibrio parahaemolyticus demonstrated by monoclonal antibodies. J. Infect. Dis., 160, 1089, 1989. 33. Honda, T., Y. Ni, and T. Miwatani. Purification of a TDHrelated hemolysin produced by a Kanagawa phenomenonnegative clinical isolate of Vibrio parahaemolyticus O6:K46. FEMS Microbiol. Lett., 57, 241, 1989.

498 34. Shirai, H. et al. Molecular epidemiologic evidence for ­association of thermostable direct hemolysin (TDH) and TDH-related hemolysin of Vibrio parahaemolyticus with gastroenteritis. Infect. Immun., 58, 3568, 1990. 35. Matsumoto, C. et al. Pandemic spread of an O3:K6 clone of Vibrio parahaemolyticus and emergence of related strains evidenced by arbitrarily primed PCR and toxRS sequence analyses. J. Clin Microbiol., 38, 578, 2000. 36. WHO. Vibrio parahaemolyticus, Japan, 1996–1998. Wkly. Epidemiol. Rec., 74, 357, 1999. 37. Taniguchi, H. et al. Cloning and expression in Escherichia coli of Vibrio parahaemolyticus thermostable direct hemolysin and thermolabile hemolysin genes. J. Bacteriol., 162, 510, 1985. 38. Bej,A.K. et al. 1999. Detection of total and ­hemolysin-­producing Vibrio parahaemolyticus in shellfish using multiplex PCR amplification of tl, tdh and trh. J. Microbiol. Methods, 36, 215, 1999. 39. Linkous, D.A. and Oliver, J.D. Pathogenesis of Vibrio vulnificus. FEMS Microbiol. Lett., 174, 207, 1999. 40. Patel, V.J., Gardner, E. and Burton, C.S. Vibrio vulnificus septicemia and leg ulcer. J. Am. Acad. Dermatol., 46, S144, 2002. 41. Ulusarac, O. and Carter, E. Varied clinical presentations of Vibrio vulnificus infections: a report of four unusual cases and review of the literature. South Med. J., 97, 163, 2004. 42. Hollis, D.G. et al. Halophilic Vibrio species isolated from blood cultures. J. Clin. Microbiol. 3, 425, 1976. 43. Paranjpye, R.N. et al. The type IV leader peptidase/N-methyltransferase of Vibrio vulnificus controls factors required for adherence to Hep-2 cells and virulence in iron-overloaded mice. Infect. Immun., 66, 5659, 1998. 44. Strom, M.S. and Paranjpye, R.N. Epidemiology and pathogenesis of Vibrio vulnificus. Microbes Infect., 2, 177, 2000. 45. Wright, A.C., Simpson, L.M. and Oliver, J.D. Role of iron in the pathogenesis of Vibrio vulnificus infections. Infect. Immun., 34, 503, 1981. 46. Hor, L.I. et al. Mechanism of high susceptibility of ironoverloaded mouse to Vibrio vulnificus infection. Microbiol. Immunol., 44: 871, 2000. 47. Hayat, U. et al. Capsular types of Vibrio vulnificus: an analysis of strains from clinical and environmental sources. J. Infect. Dis., 168, 758, 1993. 48. Huq, M.I. et al. Isolation of Vibrio-like group, EF-6, from patients with diarrhea. J. Clin. Microbiol., 11, 621, 1980. 49. Chikahira, M. and Hamada, K. Enterotoxigenic substance and other toxins produced by Vibrio fluvialis and Vibrio furnissii. Nippon Juigaku Zasshi., 50, 865, 1988. 50. Nishibuchi, M. and Seidler, R.J. Medium-dependent production of extracellular enterotoxins by non-O-1 Vibrio cholerae, Vibrio mimicus, and Vibrio fluvialis. Appl Environ Microbiol., 45, 228, 1983. 51. Jean-Jacques, W. In: Manual of Clinical Microbiology, 6th Edn., p. 465. Murray, P.R. et al. (eds.). ASM Press, Washington, DC, 1995. 52. Dalsgaard., A. et al. Clinical manifestations and molecular epidemiology of five cases of diarrhoea in children associated with Vibrio metschnikovii in Arequipa, Peru. J. Med, Microbiol., 45, 494, 1996. 53. Hlady, W.G. and Klontz, K.C. The epidemiology of Vibrio infections in Florida, 1981–1993. J. Infect. Dis., 173, 1176, 1996. 54. Rank, E.L., Smith, I.B. and Langer, M. Bacteremia caused by Vibrio hollisae. J. Clin. Microbiol., 26, 2 ,1988.

Molecular Detection of Foodborne Pathogens 55. Vollberg, C.M. and Herrara, J.L. Vibrio vulnificus infection: an important cause of septicemia in patients with cirrhosis. South Med. J., 90, 1040, 1997. 56. Zide, N., Davis, J. and Ehrenkranz, N.J. Fulminating Vibrio parahaemolyticus septicemia. Arch. Intern. Med., 133, 479, 1974. 57. Marano, N.N. et al. Stool culturing practices for Vibrio ­species at clinical laboratories in Gulf Coast states. J. Clin. Microbiol., 38, 2267, 2000. 58. Kobayashi, T. et al. A new selective isolation medium for vibrio group on a modified Nakanishi’s medium (TCBS agar medium). Jpn. J. Bacteriol., 18, 387, 1963. 59. Atlas, R.M. Handbook of Microbiological Media, p. 529. CRC Press, Boca Raton, FL, 1993. 60. Lotz, M.J. Tamplin, M.L. and Rodrick, G.E. Thiosulphate­ citrate-bile salts-sucrose agar and its selectivity for clinical and marine vibrio organisms. Ann. Clin. Lab., 13, 45, 1983. 61. Tamplin, M.L. et al. Enzyme immuno assay for identification of Vibrio vulnificus in seawater, sediment, and oysters. Appl. Environ. Microbiol. 57, 1235, 1991. 62. Cerda-Cuellar, M., Jofre, J. and Blanch, A.R. A selective medium and a specific probe for detection of Vibrio vulnificus. Appl. Environ. Microbiol., 66, 855, 2000. 63. Cerda-Cuellar, M. et al. Comparison of selective media for the detection of Vibrio vulnificus in environmental samples. J. Appl. Microbiol., 91, 322, 2001. 64. Wright, A.C. et al. Rapid identification of Vibrio vulnificus on nonselective media with an alkaline phosphataselabeled oligonucleotide probe. Appl. Environ. Microbiol., 59, 541, 1993. 65. Elliot, E.L. et al. Vibrio cholerae, V. parahaemolyticus, V. vulnificus and other Vibrio spp. In: FDA Bacteriological Analytical Manual, 8th Edn. pp. 9.01–9.27. AOAC International, Gaithersburg, MD, 1995. 66. Miceli, G.A. et al. Direct plating procedure for enumerating Vibrio vulnificus in oysters (Crassostrea virginica). Appl. Environ. Microbiol., 59, 3519, 1993. 67. McCarthy, S.A. et al. Evaluation of alkaline phosphatase-and digoxigenin-labeled probes for detection of the thermolabile hemolysin (tlh) gene of Vibrio parahaemolyticus. Lett. Appl. Microbiol., 28, 66, 1999. 68. DePaola, A. and Hwang, G.C. Effect of dilution, incubation time, and temperature of enrichment on cultural and PCR detection of Vibrio cholerae obtained from the oyster Crassostrea virginica. Mol. Cell. Probes, 9, 75, 1995. 69. DePaola, A., Motes, M.L. and McPhearson, R.M. Comparison of the APHA and elevated temperature enrichment methods for recovery of Vibrio cholerae from oysters: Collaborative study. J. Assoc. Off. Anal. Chem. 71, 584, 1988. 70. Gooch, J.A. et al. Evaluation of two direct plating methods using nonradioactive probes for enumeration of Vibrio parahaemolyticus in oysters. Appl. Environ. Microbiol., 67, 721, 2001. 71. Brasher, C.W. et al. Detection of microbial pathogens in shellfish with multiplex PCR. Curr. Microbiol., 37, 101, 1998. 72. Rivera, I.N. et al. Genotypes associated with virulence in environmental isolates of Vibrio cholerae. Appl. Environ. Microbiol., 67, 2421, 2001. 73. Rivera, I.N. et al. Method of DNA extraction and application of multiplex polymerase chain reaction to detect toxigenic Vibrio cholerae O1 and O139 from aquatic ecosystems. Environ. Microbiol., 5, 599, 2003. 74. Lyon. W.J. TaqMan PCR for detection of Vibrio cholerae O1, O139, non-O1, and non-O139 in pure cultures, raw oysters, and synthetic seawater. Appl. Environ. Microbiol., 67, 4685, 2001.

Vibrio 75. Fedio, W. et al. Rapid detection of the Vibrio cholerae ctx gene in food enrichments using real-time polymerase chain reaction. J. AOAC Int., 90, 1278, 2007. 76. Blackstone, G.M. et al. Use of a real time PCR assay for detection of the ctxA gene of Vibrio cholerae in an environmental survey of Mobile Bay. J. Microbiol. Methods, 68, 254, 2007. 77. Myers, M.L., Panicker, G. and Bej, A.K. PCR detection of a newly emerged pandemic Vibrio parahaemolyticus O3:K6 pathogen in pure cultures and seeded waters from the Gulf of Mexico. Appl. Environ. Microbiol., 69, 2194, 2003. 78. Rizvi, A.V. et al. Detection of pandemic Vibrio parahaemolyticus O3:K6 serovar in Gulf of Mexico water and shellfish using real-time PCR with TaqMan fluorescent probes. FEMS Microbiol. Lett., 262, 185, 2006. 79. Ocurra, M. et al. Genotypic analyses of Vibrio parahaemolyticus and development of a pandemic group-specific multiplex PCR assay. J. Clin. Microbiol., 41, 4676, 2003. 80. Ward, L.N. and Bej, A.K. Detection of Vibrio parahaemolyticus in shellfish by use of multiplexed real-time PCR with TaqMan fluorescent probes. Appl. Environ. Microbiol., 72, 2031, 2006. 81. Nordstrom, J.L. et al. Development of a multiplex real-time PCR assay with an internal amplification control for the detection of total and pathogenic Vibrio parahaemolyticus bacteria in oysters. Appl. Environ. Microbiol., 73, 5840, 2007. 82. Hill, W.E. et al. Polymerase chain reaction identification of Vibrio vulnificus in artificially contaminated oysters. Appl. Environ. Microbiol., 57, 707, 1991. 83. Brauns, L.A., Hudson, J.D. and Oliver, J.D. Use of the ­polymerase chain reaction in detection of culturable and nonculturable Vibrio vulnificus cells. Appl. Environ. Microbiol., 57, 2651, 1991. 84. Arias, C.R., Garay, E. and Aznar, R. Nested PCR method for rapid and sensitive detection of Vibrio vulnificus in fish, sediments, and water. Appl. Environ. Microbiol., 61, 3476, 1995. 85. Wang, H.,Fadl, A.A. and Khan, M.I. Multiplex PCR for avian pathogenic mycoplasmas. Mol. Cell. Probes, 11, 3, 1997. 86. Lee, C.Y., Panicker, G. and Bej, A.K. Detection of ­pathogenic bacteria in shellfish using multiplex PCR followed by CovaLink NH microwell plate sandwich hybridization. J. Microbiol. Methods, 53, 199, 2003. 87. Higuchi, R. et al. Simultaneous amplification and detection of specific DNA sequences. Biotechnology (NY), 10, 413, 1992. 88. Levin, R.E. The application of real-time PCR to food and agricultural systems, A review. Food Biotechnol., 18, 97, 2004. 89. Jaykus, L. Challenges to developing real-time methods to detect pathogens in foods. ASM News, 69, 341, 2003. 90. Panicker, G. et al. Detection of pathogenic Vibrio spp. in shellfish by using multiplex PCR and DNA microarrays. Appl. Environ. Microbiol., 70, 7436, 2004. 91. Campbell, M.S. and Wright, A.C. Real-time PCR analysis of Vibrio vulnificus in oysters. Appl. Environ. Microbiol., 69, 7137, 2003.

499 92. Panicker, G. and Bej, A.K. Real-time PCR detection of Vibrio vulnificus in oysters: Comparison of oligonucleotide primers and probes targeting vvhA. Appl. Environ. Microbiol., 71, 5702, 2005. 93. Oliver, J.D. The viable but nonculturable state in bacteria. J. Microbiol., 43, 93, 2005. 94. Whitesides, M.D. and Oliver, J.D. Resuscitation of Vibrio vulnificus from the viable but nonculturable state. Appl. Environ. Microbiol., 63, 1002, 1997. 95. Fischer-Le Saux, M. et al. Detection of cytotoxin-hemolysin mRNA in nonculturable populations of environmental and clinical Vibrio vulnificus strains in artificial seawater. Appl. Environ. Microbiol., 68, 5641, 2002. 96. Coutard. F., et al. Real-time reverse transcription-PCR for transcriptional expression analysis of virulence and housekeeping genes in viable but nonculturable Vibrio parahaemolyticus after recovery of culturability. Appl. Environ. Microbiol., 73, 5183, 2007. 97. Coutard., F. et al. mRNA detection by reverse transcriptionPCR for monitoring viability and potential virulence in a pathogenic strain of Vibrio parahaemolyticus in viable but nonculturable state. J. Appl. Microbiol., 98, 951, 2005. 98. Sun, F. et al. Characterization and virulence retention of viable but nonculturable Vibrio harveyi. FEMS Microbiol. Ecol., 64, 37, 2008. 99. Baffone W. et al. Retention of virulence in viable but nonculturable halophilic Vibrio spp. Int. J. Food Microbiol., 89, 31, 2003. 100. González-Escalona, N. et al. Quantitative reverse transcription polymerase chain reaction analysis of Vibrio cholerae cells entering the viable but non-culturable state and starvation in response to cold shock. Environ. Microbiol., 8, 658, 2006. 101. Kaysner, C.A. and DePaola, A., Jr. Vibrio cholerae, V. parahaemolyticus, V. vulnificus, and other Vibrio spp. In: FDA Bacteriological Analytical Manual, 8th Edn. Revision A, CFSAN, Washington, DC, 2004. 102. Ausubel FM. et al. (eds). Current Protocols in Molecular Biology, pp. 2.10–2.11. John Wiley & Sons Inc., NY, 1987. 103. Han, F. and Ge, B. Evaluation of a loop-mediated isothermal amplification assay for detecting Vibrio vulnificus in raw oysters. Foodborne Pathog. Dis., 5, 311, 2008. 104. Ren, C.H. et al. Sensitive and rapid identification of Vibrio vulnificus by loop-mediated isothermal amplification. Microbiol Res., Jun 25 [Epub ahead of print], 2008. 105. Luan, X. et al. Rapid quantitative detection of Vibrio parahaemolyticus in seafood by MPN-PCR. Curr. Microbiol., 57, 218, 2008. 106. Wright, A.C. et al. Evaluation of postharvest-processed oysters by using PCR-based most-probable-number enumeration of Vibrio vulnificus bacteria. Appl. Environ. Microbiol., 73, 7477, 2007. 107. Mertens, J. et al. Label-free detection of DNA hybridization based on hydration-induced tension in nucleic acid films. Nat. Nanotechnol., 3, 301, 2008.

36 Yersinia Mikael Skurnik

University of Helsinki and Helsinki University Central Hospital Laboratory Diagnostics

Peter Rådström Lund University

Rickard Knutsson and Bo Segerman National Veterinary Institute

Saija Hallanvuo

Finnish Food Safety Authority Eviva

Susanne Thisted Lambertz National Food Administration

Hannu Korkeala

University of Helsinki

Maria Fredriksson-Ahomaa Ludwig-Maximilian University

Contents 36.1 Introduction.................................................................................................................................................................... 502 36.1.1 Taxonomical Aspects of Genus Yersinia.......................................................................................................... 502 36.1.2 Y. pestis—Causative Agent of Plague.............................................................................................................. 502 36.1.2.1 Biology and Pathogenesis................................................................................................................ 502 36.1.2.2 Epidemiology................................................................................................................................... 503 36.1.2.3 Conventional and Molecular Diagnosis........................................................................................... 503 36.1.3 Y. pseudotuberculosis....................................................................................................................................... 503 36.1.3.1 Biology and Pathogenesis................................................................................................................ 503 36.1.3.2 Epidemiology................................................................................................................................... 505 36.1.3.3 Conventional and Molecular Diagnosis........................................................................................... 505 36.1.4 Y. enterocolitica................................................................................................................................................ 505 36.1.4.1 Biology and Pathogenesis................................................................................................................ 505 36.1.4.2 Epidemiology................................................................................................................................... 506 36.1.4.3 Conventional and Molecular Diagnosis........................................................................................... 507 36.2 Methods.......................................................................................................................................................................... 508 36.2.1 Reagents and Equipment ................................................................................................................................. 508 36.2.2 Food and Environmental Sample Collection and Preparation......................................................................... 509 36.2.3 Detection Procedures.........................................................................................................................................510 36.2.4 Quality Assurance/Aspects When Using PCR..................................................................................................511 36.3 Conclusions and Future Perspectives..............................................................................................................................513 Acknowledgments.......................................................................................................................................................................513 References...................................................................................................................................................................................513 501

502

Molecular Detection of Foodborne Pathogens

36.1  Introduction In this chapter we will introduce the bacteria of genus Yersinia, describe their taxonomical, biological, patho­ logical, and epidemiological properties and discuss molecular methods associated with their detection, isolation, and characterization.

36.1.1 Taxonomical Aspects of Genus Yersinia The genus Yersinia—a member of family Enterobacteriaceae of the γ-purple bacteria1 now comprises 14 species, three of which contain human pathogenic strains: Y. pestis, Y. pseudotuberculosis, and Y. enterocolitica. Y. pestis is the causative agent of the feared bubonic plague that spreads from infected rodents to humans via a flea vector.2 Pathogenic strains of Y. pseudotuberculosis, and Y. enterocolitica cause yersiniosis which usually is a mild diarrheal disease, and the infection usually takes place by ingestion of contaminated food stuffs. Y. pseudotuberculosis, and Y. enterocolitica and related species (Y. aldovae, Y. aleksiciae, Y. bercovieri, Y. frederiksenii, Y. intermedia, Y. kristenseni, Y. mollareti, Y. rohdei, Y. massiliensis, and Y. similis, the two latter of which were only recently described3,4) have several serovariants whereas Y. pestis has three biovariants but is serologically of a single type. Five serotypes have been described for Y. ruckeri that causes the enteric red mouth disease of fish.5 A

In Figure 36.1 the Yersinia strains for which the complete genomic sequences are available are used to illustrate the phylogenetic relationships between Yersinia species and in particular individual Y. pestis strains. The close relation between Y. pestis and Y. pseudotuberculosis can be seen in Figure 36.1A and the relation within the relatively well conserved Y. pestis group is shown in Figure 36.1B. Y. pestis is considered a recently (within the last 1,500–40,000 years) diverged clone of Y. pseudotuberculosis serotype O:1b.6–9 Although Y. pestis is not usually considered as a food pathogen we will include it also here as there is a potential threat that it could be used as a biological weapon by food contamination. Indeed, eating contaminated camel meat has caused plague cases.10,11

36.1.2  Y. pestis—Causative Agent of Plague 36.1.2.1 Biology and Pathogenesis Among the all pathogenic Yersiniae, Y. pestis causes the most fatal and devastating disease—bubonic plague, that was during the Middle Ages known as a Black Death. The number of people killed by this bacterium in world history approaches 200 million. It has been responsible for three human pandemics—the Justinian plague in the fifth to seventh century, the Black Death in the thirteenth to fifteenth century, which is claimed to have killed one-third of the Yersinia pestis (16 strains) Yersinia pseudotuberculosis (2 strains) (IP 32953 IP 31758) Yersinia bercovieri (ATCC 43970) Yersinia mollaretii (ATCC 43969) Yersinia enterocolitica (subsp. enterocolitica) Yersinia frederiksenii (ATCC 33641) Yersinia intermedia (ATCC 29909)

B

KIM (Mediaevalis) K1973002 (Mediaevalis) Nepal516 (Antiqua) B42003004 (Antiqua) 91001 (Mediaevalis/Microtus) UG05-0454 (Antiqua) Antiqua (Antiqua) Pestoides F (Antiqua) E1979001 (Antiqua) IP275 (Orientalis) CA88-4125 (Orientalis) F1991016 (Orientalis) CO92 (Orientalis)

Angola (Antiqua)

FV-1 (Orientalis)

MG05-1020 (Orientalis)

Figure 36.1  Whole genome comparison of Yersinia strains whose genome sizes are approximately 4.5–4.8 Mb. The dendrograms are based on a distance matrix created from average blast score values of 5.4 million cross-blast comparisons spanning the entire genomes. Panel A: relation at genus level. Panel B: relation between Y. pestis genomes. Biovariant classifications are indicated. The biovariants might not be completely monophyletic, which has also been observed with multilocus variable number of tandem repeat analysis (MLVA).6

503

Yersinia

European population and modern plagues since the 1870s. Although plague is no longer a serious threat it still occurs occasionally in developing countries.2,12 Y. pestis is a vector-borne pathogen, and has a complex life-cycle involving mammalian reservoir (primarily rodents) and a flea vector. The flea acquires the bacteria by taking a blood meal from an infected mammal. In the flea gut, the bacterium replicates and forms a clumped mass, which blocks the flea intestine from receiving further blood meals. An infected insect is, in effect, starving and actively looking for a new host. When an infected flea bites a human, it regurgitates fragments of mass containing microbial pathogen. The bacteria disseminate rapidly to the lymphatic system and locate in the nearby lymph nodes, where the massive inflammatory response causes painful swellings, the factual buboes which are the hallmark symptoms of bubonic plague. At this point, Y. pestis may enter the bloodstream and travel to the lungs, where it can cause pneumonic plague. Pneumonic plague can be transmitted by aerosols from one human to another. Ninety percent of untreated victims infected by this route die within a few days due to septic shock.12 Wild type Y. pestis strains carry three virulence ­factor encoding plasmids: pPst, pFra, and pYV (also known as pPCP1, pMT1, and pCD1, respectively). While the latter, encoding the type III secretion system (T3SS) and the secreted effector toxins called Yops, is also present in the other pathogenic Yersiniae, the other two plasmids are unique to Y. pestis. The 9.6 kb pPst plasmid carries the pla gene encoding plasminogen activator, Pla, a very important protease indispensable for bacterial dissemination after subcutaneous injection into mammalian tissues.13 Other Y. pestis unique factors, like capsular protein F1 and Ymt (Yersinia murine toxin), are encoded by the genes present on pFra plasmid. They appear to prevent phagocytosis and play a role in colonization of the flea gut.14 36.1.2.2 Epidemiology Serologically Y. pestis strains are very homogenous because their lipopolysaccharide (LPS) is rough, i.e., lacking O-antigen.15 Y. pestis has three biovariants: mediaevalis, orientalis, and antiqua, based on differences in fermentation of glycerol (+, −, +) and ability to reduce nitrate to nitrite (−, +, +), respectively.16,17 Variety orientalis is endemic in India, southwest Asia, South America and western North America; antiqua is present in southeast and northern Asia (China and Russia), and Africa; and mediaevalis is limited to Turkey and Iran.17 During the relatively short time Y. pestis evolved from the less virulent Y. pseudotuberculosis the pre-Y. pestis bacterium experienced very significant genetic changes that collectively have resulted in the huge increase in virulence when compared to Y. pseudotuberculosis. 36.1.2.3  Conventional and Molecular Diagnosis The samples from where Y. pestis are conventionally cultured are sputum or bronchial washes, blood, aspirates or biopsies, but also from environmental or other nonclinical

samples. The CDC Plague home page (http://www.cdc. gov/ncidod/dvbid/plague/index.htm) informs that suspicion for the ­presence of Y. pestis is raised if Gram-negative and/ or bipolar-staining coccobacilli are seen on a smear taken from affected tissues after Gram or other differential staining. A presumptive identification of Y. pestis can be made if the capsule F1 antigen can be detected from the specimen by immunofluorescence. Y. pestis grows slowly and usually visible colonies appear in 48 h. The potential Y. pestis colonies can be identified as Y. pestis by biochemical tests and use of specific antisera and bacteriophages. In the present time, PCR and other sequence-based identification methods are used to confirm the identification (see below). Due to increased funding to research biothreat agents, recent years have evidenced a burst of molecular methods designed for the detection of Y. pestis. The methods range from simple single-PCR assays via multiplex real-time PCR assays to microarray-based protocols and also include other detection technologies (Table 36.1). Specific attention has been placed on the danger that the very close relative Y. pseudotuberculosis poses on the detection method as a false-positive. Handling of Y. pestis strains requires Biosafety Level 3 (BSL-3) capabilities for appropriate safety. The reason is that the most important route of Y. pestis exposure of laboratory workers is by inhalation. BSL-3 is used for work with indigenous agents when the potential for infection by aerosol is real and the disease may have serious or lethal consequences.18 Special engineering design criteria for BSL-3 laboratories for laboratory work on Y. pestis involve the access zone, sealed penetrations, and directional airflow to aid in protecting the employee from respiratory exposure. In addition appropriate cabinets; dress codes, risk assessments, deactivation, and emergency procedures are needed. For deactivation of Y. pestis 10% sodium hypochlorite can be used before autoclaving and formaldehyde fumigation can be used for cabinet and room decontamination. Consequently, the isolation and molecular detection methods of Y. pestis have to be implemented in order to fulfil the biosafety requirements.

36.1.3  Y. pseudotuberculosis 36.1.3.1 Biology and Pathogenesis Y. pseudotuberculosis is an enteropathogen widely spread in the environment (soil, water, vegetables). It was first described in 1884.19 This bacterium is a primary pathogen of wild and domestic animals in all continents.20 In humans, Y. pseudotuberculosis infections are not frequent although outbreaks associated with consumption of water or food stuffs contaminated with animal faeces are reported. Symptoms of infection vary from abdominal pain and fever to septicemia and mesenteric lymphadenitis. In most cases, the infection is self-limiting and can be effectively treated with antibiotic therapy.21–23 Major virulence factors associated with Y. pseudotuberculosis are the pYV-encoded Yop proteins that enter host mammalian cells through T3SS.24 Other virulence factors are

504

Molecular Detection of Foodborne Pathogens

Table 36.1 Molecular Methods for Detection or Identification of Y. pestis Method Conventional single PCR assays

Target Sequence Chromosome or Plasmid pla 16S rDNA, caf1 caf1, pla, lcrV, 16S rDNA 41.7 kb insertion site caf1 16S and 23S rDNA pla 28 signature genes

Real-time PCR assays

wzx-wbyIJ pla gyrA pla Not given Chromosomal YP48 gene pla, pim, caf1 pst pla 16S rDNA

Multiplex PCR assays

caf1, pla, lcrV, irp2 caf1, pla, lcrV, ymt, 16S rDNA caf1, pla, yopM, inv yihN, caf1, pla, lcrV Not given entF3, caf1, pla, pCD1-region pla, ypo2088, wzz

Microarray-based assays

Multiple targets Multiple targets Not given

Other methods

F1-antigen F1-antigen F1-antigen F1-antigen

Not given

Samples, Detection Method and Comments

Reference

Wild fleas. Gel detection. Soil samples. Gel detection and sequencing. Suitable for identification of Yersinia species. Gel detection. The target is specific for Y. pestis. Gel detection. Madagaskar field study. Gel detection. Gel detection and a fluorescent probe for in situ hybridization. 6th century teeth. Gel detection and sequencing. Genes identified by microarray. PCR analysis by gel detection. Specific identification of Y. pestis. Gel detection. Spiked fleas, blood oropharyngeal swabs. Ciprofloxacin resistance. Sputum. Field-deployable RAPID (“Ruggedized” advanced pathogen identification device). Differentiation between Y. pestis and Y. pseudotuberculosis. Detection by TaqMan minor groove binder probe. Detection by dual-labeled fluorogenic probe. Detection by TaqMan fluorogenic probe. Simultaneous detection of other biothreat agents. Detection by TaqMan minor groove binder probe. Detection by TaqMan probe.

129 130 131 132 133 134 135 136 55 137 138, 139 115 140 116 141 142 143 144

Samples from 1986 Paraiba outbreak. Gel detection. Real time PCR. Detection using fluorescent probes.

145 146

Gel detection. Detection by TaqMan probes. Rapid simultaneos detection of select category A bacterial agents from blood. Detection by SYBR Green and Tm. Real-time multiplex PCR. Detection by TaqMan probes. Real-time multiplex PCR. Dual-labeled fluorogenic probes. Plasmid and chromosomal targets. wzz detects both Y. pestis and Y. pseudotuberculosis. Internal ampification control. Multiplex PCR microarray. Simultaneous detection of bioterror agents in blood. Multiplexed high-density DNA array. Multiplexed PCR-coupled liquid bead array for simultaneous detection of many biothreat agents. Rapid diagnostic test (RDT) to study a medieval burial place. Fiber optic biosensor for detection from blood etc. Magnetic-beads based magnetic biosensor for detection from serum and blood samples. Immunochromatography, Plague BioThreat Alert test strips, ABICAP columns, ELISA, flow cytometry, and IF-microscopy. Immunoassay plus real-time PCR to analyze aerosols.

147 117 148

adhesion molecules, invasin and YadA25 and the superantigenic toxin designated YPM (Y. pseudotuberculosis-derived mitogen) which mediates activation of the immune system.26 In addition, the LPS O-polysaccharide27,28 and type IV pili29 contribute to pathogenicity.

119 118

149 150 151 152 153 154 155

156, 157

The current Y. pseudotuberculosis antigenic scheme lists 21 O-serotypes.30,31 The division into different groups is based on different O-factors that reflect the different antigenic sugar residues present in the O-polysaccharide (O-antigen).31 Some of the O-factors (specifically those carrying 3,6-dideoxyhexoses)

505

Yersinia

are shared with the O-antigenic structures of Salmonella and E. coli and may cause cross-reactivity.32,33 36.1.3.2 Epidemiology Y. pseudotuberculosis infections among humans have been reported from all continents, but the incidence appears to be below that of Y. enterocolitica. Epidemiologically there are some differences in the prevalence of the serotypes in human isolates. In Europe, most often the isolates belong to serotypes O:1–O:334 while to serotypes O:4b, O:3, O:5a, and O:5b in Japan.35 In recent years, several outbreaks have been reported in Finland, Russia, and Japan.35–39 The outbreaks in Finland have involved large-scale kitchens and mainly affected children in day-care centers and schools.39 In culture-based detection surveys the isolation rates from different animal sources are low; 0.1–4%.40–49 The principal reservoirs for Y. pseudotuberculosis are believed to be rodents, wild birds and domestic animals (especially pigs and ruminants).50 In Japan, Y. pseudotuberculosis strains isolated from monkeys, goats, rabbits, and guinea pigs showed disease symptoms while cattle, swine, dogs, cats, hares, and rats were considered as carriers.35 The healthy carriers may get ill and excrete the bacteria if exposed to a stress like cold and humid weather or starvation. Y. pseudotuberculosis has been isolated sporadically from the oral cavities and caecal contents of asymptomatic finishing pigs.43,51 The environment can easily be contaminated by the feces of infected animals, mainly wild animals like deer, rodents, and birds. Y. pseudotuberculosis in soil, water or vegetables can survive for a long time as in rivers, wells or mountain streams in Japan and Korea.35,50 Y. pseudotuberculosis has been sporadically isolated from fresh food products in Finland and Russia or from pork in Japan.37,39,52 The food products may get contaminated with Y. pseudotuberculosis during irrigation, harvesting, packing, shipping or processing. Some recent epidemiologic investigations in Finland have linked outbreaks of Y. pseudotuberculosis O:1 and O:3 infections to domestically grown iceberg lettuce and carrots.36,38,39 In the outbreak linked to carrots, washing and peeling equipment was shown to be contaminated with the outbreak strain, however, the exact mechanism of contamination of carrots in the farm could not be clarified.38 A combination of direct contact with wildlife feces during the storage and cross-contamination of the equipment were the most likely contributing factors. In another outbreak, iceberg lettuce was probably contaminated with irrigation water.36 36.1.3.3  Conventional and Molecular Diagnosis The isolation of Y. pseudotuberculosis is demanding, especially when food and environmental samples are studied since Y. pseudotuberculosis grows slowly and the minute colonies are easily overgrown by other bacteria. In addition, only limited information is available on reliable protocols of Y. pseudotuberculosis recovery from foods and environment. Cold enrichment for two weeks in slightly selective ­phosphate-buffered saline broth supplemented with 1%

mannitol and 0.15% bile salts and an alkali treatment after cold enrichment just before plating on solid media was shown to improve the isolation rate of Y. pseudotuberculosis.43 Selective agar plates are needed for isolation of Y. pseudotuberculosis strains. However, the growth of Y. pseudotuberculosis is often poor and delayed as many selective agars are designed for Y. enterocolitica. The Yersinia-selective Cefsulodin-irgasan-novabiocin (CIN)-agar is the most commonly used agar also for Y. pseudotuberculosis,53 however, some Y. pseudotuberculosis strains may not grow and the use of a second agar such as McConkey agar is advised despite its low selectivity.54 Y. pseudotuberculosis can be identified using some biochemical key tests or with commercial identification kits. Among Y. pseudotuberculosis strains there is little variation in biochemical reactions, except for the sugars melibiose and raffinose, and salicin, which can be used to divide the species into four biotypes.30 The commercially available anti-O:1–O:6 antisera have been used for serotyping of Y. pseudotuberculosis, however, no antisera are commercially available to other serotypes. A PCR-based O-genotyping scheme that covers all the 21 O-serotypes was recently published.55,56 The majority of Y. pseudotuberculosis strains are pathogenic; however, a number of phenotypic characteristics associated with the virulence are available to evaluate the pathogenic potential of Y. pseudotuberculosis isolates, e.g., PCR-methods specific for virulence factor encoding genes.20 The isolation of pathogenic Yersinia from asymptomatic carriers, food and environmental samples is always demanding. The low isolation rates of Y. pseudotuberculosis in naturally contaminated samples may be due to the limited sensitivity of culture methods. Some PCR methods have been reported to identify and characterize Y. pseudotuberculosis strains (Table 36.2). However, most of them have been designed to study pure cultures. A wider application of PCR assays would probably provide better estimations of the occurrence of Y. pseudotuberculosis in animal, food, and, environmental samples.

36.1.4  Y. enterocolitica 36.1.4.1 Biology and Pathogenesis Y. enterocolitica was first isolated in the 1930s57 and initially described in 1963 as Pasteurella X.58,59 Y. enterocolitica are facultative anaerobic rods, do not possess a capsule and are motile below 28°C. Y. enterocolitica is able to grow at temperatures ranging from 4°C to 43°C. However optimal growth and metabolic activity for this species is 28–30°C. Ability to grow at refrigeration temperatures allows it to multiply in stored foods.60 Y. enterocolitica and related species comprise over 70 serotypes, which are mainly determined by the variability of O-antigen.61 The human and animal pathogenic Y. enterocolitica strains carry the 70–75 kb virulence plasmid of Yersinia, pYV, and most commonly belong to serotypes O:3, O:5,27, O:8 and O:9.62 Less frequently encountered pathogenic serotypes are O:1,2,3, O:4,32, O:13a,18, and O:21.

506

Molecular Detection of Foodborne Pathogens

Table 36.2 PCR-Based Detection Methods for Y. pseudotuberculosis Method

Target Sequence Chromosome or Plasmid

Comments on the Samples and Detection Method

Single PCR assays

inv, virF(lcrF)a

Pure culture and gel detection.

158

Pure culture, gel detection and in situ hybridization.

134

16S and 23S rDNA

Multiplex PCR assays

b

wzzc

Chromosomal target part of O-antigen gene cluster.

yadA ailc

Stool samples, real-time detection using SYBRGreen. Enrichment, DNA extration. Includes IAC. Taqman probe. Specific for most serotypes. Pure culture and gel detection. Food samples and gel detection. Nested multiplex PCR and gel detection. Pure culture and gel detection. Blood samples, nested multiplex PCR and gel detection. Pure culture and gel detection; useful for O-genotyping.

inv, virF(lcrF)a inv, virF(lcrF)a inv, virF(lcrF)a aila, virF(lcrF)a 16S rDNAb fcl, prt, manB, abe, wbyL, wbyH, ddhAB, wbyK, wzx

Reference

55 159 121 160 161 162 106 149 55

Both Y. pseudotuberculosis and Y. enterocolitica will be detected. Capable to differentiate between Y. pestis, Y. pseudotuberculosis and Y. enterocolitica. c Both Y. pseudotuberculosis and Y. pestis will be detected. a

b

As a species Y. enterocolitica comprises organisms that differ in biochemical and genetic characteristics and can be classified into six biotypes known as 1A, 1B, 2, 3, 4, and 5.63 These, on the other hand, fall into three lineages with different pathogenic potential: (i) biotype 1A strains that are nonpathogenic and lack pYV, (ii) biotype 2–5 strains that carry pYV but are weakly pathogenic and do not kill mice, and (iii) biotype 1B strains that carry pYV and are mouse-lethal and highly pathogenic. The biotype 1A strains are usually called nonpathogenic or environmental; however, they are often isolated also from stool samples of healthy humans and animals. Several chromosome and pYV-encoded virulence factors contribute to the virulence of Y. enterocolitica and many of them are common with Y. pseudotuberculosis and Y. pestis. The pYV encodes an outer membrane protein YadA (Yersinia adhesin A), the T3SS and the effector Yops.64–69 YadA is an outer membrane protein, which is an important virulence factor of Y. enterocolitica, but seems to be of less significance for the virulence of Y. pseudotuberculosis,67,70–73 and in Y. pestis YadA is not expressed as yadA is a pseudogene.69,74 At least two chromosomal encoded virulence factors, Inv (invasion) and Ail (attachment invasion locus), mediate cell invasion.75–79 The chromosome encoded Yst (Yersinia stable toxin) of Y. enterocolitica resembles closely the heat-stable toxin of enterotoxigenic Escherichia coli, however, the role of Yst in the pathogenesis of Y. enterocolitica diarrhea in humans is at present uncertain.80 Similar to Y. pseudotuberculosis also LPS contributes to the pathogenicity of Y. enterocolitica.72,81–87 The major difference in virulence between the biotype 1B and other pathogenic strains is presence of the iron uptake system in biotype 1B strains. Thus biotype 1B serotype O:8 strains kill mice while biotype 4 serotype O:3 strains do not unless iron is made available to bacteria by pre-treatment of mice with desferroxamine or iron.88

Y. enterocolitica infection can cause a variety of symptoms depending on the age of the person infected.89–91 Common symptoms in young children are fever, abdominal pain and diarrhea. Diarrhea is a more dominant symptom in Y. enterocolitica infections when abdominal pain and fever is more frequent in Y. pseudotuberculosis infection. In older children, right-sided abdominal pain and fever can be the predominant symptoms and may be confused with appendicitis. Occasionally, typically among adults, complications such as joint pains (reactive arthritis) and skin rash (erythema nodosum) may occur. Antimicrobial therapy is not considered essential in the treatment of uncomplicated enterocolitis. In more severe infections, like systemic infection and bacteremia, antimicrobials may be useful. 36.1.4.2 Epidemiology Y. enterocolitica has been isolated from humans on all continents.90 Since 2004, several member states in European Union are reporting the incidences of Y. enterocolitica infections. It is still the third most frequently reported zoonosis in Europe, even though there seems to be an overall decreasing trend over the last years.92 In 2006, the highest incidences of 15.1, 12.1, 6.3 and 6.2 cases per 100,000 inhabitants were observed in Finland, Lithuania, Germany, and Sweden, respectively. In Austria, the incidence was only 1.9, however, seroprevalence of anti-Yop antibodies among 750 healthy Austrians was shown to be 29.7%, which indicates that the majority of infections is either subclinical or mild.93 The majority yersiniosis cases are sporadic and domestically acquired.92 No seasonal distribution has been reported in Europe. The most yersiniosis cases in Europe have been reported among children in the 0–4 and 5–14 year age groups representing 32% and 20% of cases, respectively. Yersiniosis is rare in US (www.cdc.gov/ ncidod/dbmd/diseaseinfo/yersinia), with an incidence of one case per 100,000 people.

Yersinia

Pigs are mostly asymptomatic carriers of human pathogenic strains, in particular strains of bioserotype 4/O:3, also sporadically serotype 5,27, O:9 and O:8 strains have been isolated from pigs. Y. enterocolitica O:9 strains have been isolated from dogs and cats and, in addition, from stools of cattle and goats in infected herds in France.94 In Japan, serotypes O:3, O:8 and O:9 have been isolated from wild rodents, especially from field mice.95 Strains of very rare pathogenic bioserotypes, such as bioserotype 5/O:2,3, which has been isolated from sheep, hares, and goats, and bioserotype 3/O:1,2a,3 from chinchillas. Most Y. enterocolitica isolates recovered from environmental samples are nonpathogenic, however, bioserotype 4/O:3 strains have occasionally been isolated from pig slaughterhouses and butchers.96 Pathogenic Y. enterocolitica strains are frequently isolated from slaughtered pigs, especially from tonsils but also from submaxillar lymph nodes, the intestine, and feces.96 A close genetic relationship between pig and human Y. enterocolitica strains has been demonstrated by several DNAbased methods.62 Also the surfaces of freshly slaughtered pig carcasses are a good source of Y. enterocolitica due to spread of the bacterium via tonsils and intestinal content during the slaughter and dressing operations.97 The most common transmission route of this pathogen is thought to be via contaminated food especially pork and pork products. The preparation of raw pork intestine (chitterlings) has shown to be particularly risky.98 Two case-control studies have shown that yersiniosis is associated to consumption of raw or undercooked pork products and untreated water.99,100 Direct transmission from pigs to humans is possible. Slaughterhouse workers and pig farmers in Finland have shown to have elevated antibody levels to Y. enterocolitica O:3 twice so frequently as grain and berry farmers.101 Stream water may be one vehicle to transmit serotype O:8 from wild rodents to humans in Japan.95 Pet animals have also been suspected of being sources for human infections because of their close contact with humans, especially young children. Direct person-to-person contact has not been demonstrated, but Lee et al. reported Y. enterocolitica O:3 infections in infants who were probably exposed to infection by their caretakers.102 Indirect person-to-person transmission has apparently occurred in several instances by transfusion of blood products.103 36.1.4.3  Conventional and Molecular Diagnosis Several different cultural methods for isolation of Y. enterocolitica from food have been described.104 So far no single isolation method is available for the recovery of all bioserotypes. The culture methods usually involve an enrichment step in one or two enrichment broths, plating onto one or two selective agar plates, identification of typical colonies, biotyping, serotyping, and testing for virulence properties. Pathogenic Y. enterocolitica strains have been detected from different natural sources, such as animals, food, and environment both by culture and by PCR. The detection efficiency of Y. enterocolitica in nonhuman sources is clearly lower with

507

culture compared to PCR (Table 36.3). The ­isolation rate of this pathogen is low even from raw pork, except for pig tongues and edible offal (heart, liver, and kidneys).105 Pathogenic Y. enterocolitica strains have not been isolated from salami and heat-treated meat products containing pork106,107 and only a few times from beef, poultry, and milk samples.104 In these cases, cross-contamination from pork has probably occurred during processing, packing or handling. Cold enrichment in different solutions has been widely used for naturally contaminated samples. The benefit of this method is that all pathogenic strains can multiply in theses broths. The major disadvantages are the presence of nonpathogenic Yersinia and other psychrotrophic bacteria, which also multiply during enrichment, and the long incubation period, typically 21 days. By treating cold enrichments with potassium hydroxide (KOH) the background flora can sometimes be reduced, making selection of Yersinia colonies less laborious. Some selective broths for isolation of pathogenic Y. enterocolitica at higher temperatures have been developed to increase the selectivity and to shorten the incubation time.104 The most often used media are irgasan-ticarcillin­potassium chlorate (ITC) broth and modified Rappaport broth (MRB), which have been designed for recovery of strains of bioserotype 4/O:3. Both media have shown to inhibit strains of bioserotypes 2/O:5,27, 2/O:9 and 1B/O:8. Enrichment in ITC is included in the method of the International Standard Organization (ISO 10273:2003), which is the mostly used method for food samples in Europe. Many different selective agar plating media are available for isolation of Y. enterocolitica from naturally contaminated samples. The most commonly used selective agar plates are CIN, Salmonella-Shigella deoxycholate calcium chloride (SSDC) and MacConkey (MAC) agar plates. CIN agar is the most frequently used agar for naturally contaminated samples because of the high confirmation rate of presumptive isolates and its relatively high selectivity for fecal samples. CIN and SSDC agar plates are included in the ISO (10273:2003) method. Presumptive urea-positive Yersinia isolates can be identified using selected biochemical key tests or by commercial identification tests. Biotyping of Y. enterocolitica isolates is recommended because the biotype is a reliable marker of pathogenicity.92 At least five different biochemical tests are needed to distinguish the six biotypes.63 As already mentioned isolates belonging to biotype 1A are regarded as nonpathogenic. Serotyping using commercial O:3, O:5, O:8 and O:9 antisera has been used extensively, however, these antigens can sometimes be found in nonpathogenic Y. enterocolitica strains and in other Yersinia species. Finally, it is very important to asses the pathogenic potential of the isolates because the majority of strains recovered from asymptomatic carriers, food, and environmental samples are nonpathogenic and have no clinical significance. In addition to using virulence-factor-gene-specific PCR, the pyrazinamidase test and a number of phenotypic characteristics associated with the virulence plasmid (calcium dependence, detected as growth restriction on magnesium oxalate

508

Molecular Detection of Foodborne Pathogens

Table 36.3 Detection of Pathogenic Y. enterocolitica by Culture and PCR No. of Samples Sample

Culture Positive

PCR Positive

Animal 185 252 212 255 2793 257 518 98

48 0 72 0 114 0 0 0

58 90 186 80 345 103 81 9

Tofu

Finland Finland USA Finland USA Sweden Norway Sweden Sweden Finland Finland Finland Norway USA

Food 51 34 350 255 350 100 300 91 97 43 200 101 200 50

40 17 8 4 0 5 6 6 0 0 0 0 0 0

47 21 278 63 133 35 50 9 11 0 0 3 6 6

Water Slaughterhouse

Canada Finland

Environmental 105 89

1 5

11 12

Pig tonsils

Pig feces Pig mesenteric lymph node Pig carcass Poultry carcass

Pig tongues Pig offal Chitterling Minced pork

Pork Salami Chicken Fish Lettuce

Country

Finland USA Switzerland USA USA USA Sweden Sweden

Total

Source: This table has been compiled from the reports of Lindblad, M. et al., J. Food Prot., 70, 1790, 2007; Lambertz, S. T. and Danielsson-Tham, M.L., Appl. Environ. Microb., 71, 3674, 2005; Lambertz, S. T. et al., J. Food Prot., 70, 335, 2007; Fredriksson-Ahomaa, M. and Korkeala, H., Adv. Exp. Med. Biol., 529, 295, 2003; Bhaduri, S., Wesley, I.V. and Bush, E.J., Appl. Environ. Microb., 71, 7117, 2005; Lindblad, M. et al., J. Food Prot., 69, 2875, 2006.

agar, autoagglutination at 35–37°C, and uptake of Congo red and crystal violet) could be applied.104 It has recently been claimed that the development of a simple, sensitive, specific and rapid identification system applicable to the diagnosis of Y. enterocolitica is a challenge for the future. The isolation of Y. enterocolitica has been associated with difficulties and it has been suggested that the lack of appropriate analysis methods may mask the occurrence of the bacterium leading to underestimation of the true prevalence. Reasons for the difficulties encountered may include the sensitivity of the bacterium to high concentrations of background flora present in food samples, but also the heterogeneity of the species. It has been indicated that pathogenic Y. enterocolitica can be detected more rapidly and with better specificity through the use of DNA-based methods, especially PCR. It has been shown that PCR-based methods indicate a higher prevalence of pathogenic Y. enterocolitica

than traditional methods (Table 36.3). Table 36.4 summarizes some PCR-based detection methods available for Y. enterocolitica.

36.2  Methods 36.2.1 Reagents and Equipment The following equipment and materials are needed to carry out the real-time PCR-based detection method: a real-time PCR system such as the 7500 Real-time PCR system instrument (Applied Biosystems, Foster City, CA), Stomacher and Stomacher bags with filter, heating block, microcentrifuge, automatic micropipettes and filtertips, incubators for 25°C and 30°C, 96-well PCR plates or eight-well PCR strips, 0.5 ml and 1.5 ml microtubes. The reagents are listed within the description of the protocols.

509

Yersinia

Table 36.4 Molecular Detection Methods Available for Y. enterocolitica Methodology Single PCR

Target Sequence Chromosome or Plasmid yadA lcrE wbcT (rfbB)

Multiplex PCR assay

ail, virF bipA, ERIC ail, virF yadA, 16S rDNA ail, virF

Real-time PCR

yadA per 16S rDNA ail

PCR-DNA microarray

Other methods

ail yst virF, ail, yst, blaA, 16S rDNA 23S rDNA gyrB rpoB 16S rRNA Surface antigens rRNA O:3 LPS

Sample Preparation and Comments

Reference

Enrichment, nested PCR. Gel detection. DNA extraction. Gel detection. Detects also Y. pseudotuberculosis and Y. pestis. Gel detection. Specific for serotype O:3. Enrichment. Two nested single PCRs. Gel detection. Specific for all pathogenic serotypes. DNA extraction. Real-time analysis. Enrichment/DNA extraction method evaluation. Gel detection. Enrichment/buoyant density gradient centrifugation and Yersinia PCR-compatible enrichment (YPCE) medium. Gel detection. Enrichment. Most probable number. Multiplex PCR for quantitation. Gel detection. Enrichment/DNA extraction. Detection by SYBR Green. DNA extraction. Includes IAC. Detection by TaqMan probe. Specific for serotype O:9. Flotation to enrich target bacteria. Detection by SYBR Green. Enrichment, DNA extraction. Includes IAC. Taqman probe. Specific for all pathogenic serotypes. Enrichment. DNA extraction. TaqMan based. Enrichment. DNA extraction. TaqMan-based. DNA extraction. Multiplex PCR. Detection by microarray. Enrichment. Analysis of PCR products by oligonucleotide array. DNA extraction. Analysis of PCR products by oligonucleotide array. DNA Extraction. PCR-DGGE (denaturing gradient gel electrophoresis). RNA Extraction. Microfluidic chip-based system. Enrichment. Multiplexed chemiluminescent immunoassay. Cell lysis. RNA Hybridization-electrochemiluminescence. Enrichment. Surface adhesion immunofluorescence.

163 164

36.2.2 Food and Environmental Sample Collection and Preparation No analysis will be accurate unless the sample withdrawn is representative of the bulk matrix. The choice of sampling technique depends on the aim of the analysis and the practical requirements to ensure the procedure that most accurately reflects the true microbiological status of the clinical or food sample. The isolation of Y. enterocolitica and Y. pseudotuberculosis has been associated with difficulties mainly due to overgrowth of the slower-growing Yersinia bacteria by the abundant background flora. Consequently, the sampling of Yersinia is often very complex due to the distribution of the bacteria in the sample, but also due to the dynamic and heterogeneous properties of the sample matrix. In a review by Fleet108 on microorganisms in complex ecosystems, several factors have been highlighted for microbial analysis, such as diversity of population, microbial viability, changes in population, spatial distribution of the species in the sample, etc. All these factors should also be considered when a sampling technique for PCR detection is chosen. Sampling plans are available for some foodborne pathogens, but not for pathogenic Y. enterocolitica.109 Furthermore, the sample

165 166 167 168 169, 170 171 159 172 173 123 174 175 176 177 178 179 180 181 182 183

transportation to the laboratory may also affect its quality and must be taken into account.110 A guidance document under regulation (EC) No 882/2004 provides information concerning microbiological sampling and transportation of food samples and also summarizes international recognized standards and guides for sampling.111 The next step after sampling is sample preparation. The objectives of this step prior to diagnostic PCR are: (i) to reduce the size of the heterogeneous bulk sample to a small homogeneous PCR sample in order to ensure negligible variations between repeated sampling, (ii) to concentrate the target organism to a concentration within the practical operating range of the PCR assay, and (iii) to remove or neutralize substances that may interfere with the PCR amplification. Many sample preparation methods are laborious, expensive, and time consuming, or do not provide the desired template quality. The most common approach is to extract nucleic acids directly from the sample matrix in order to generate a homogeneous sample of high template quality. The sample, free from most PCR inhibitors, may be stored in an appropriate buffer, such as Tris-EDTA (TE) buffer. The drawback with extracting DNA directly from the sample is the poor detection limit due to small volumes and high amounts

510

of nontemplate DNA. Therefore, a number of ­sample ­preparation methods prior to DNA extraction have been developed.112 For example, in order to detect low concentrations of pathogens in food an enrichment cultivation step is commonly employed. The aim of the enrichment culture is to provide detectable concentrations of viable target cells prior to PCR. Furthermore, enrichment cultures prior to PCR analysis serves additional purposes, including the dilution of PCR-inhibitory substances present in the sample matrix, dilution of dead target cells and last, but not least, the possibility to isolate viable cells. For example, Knutsson et al.113 constructed a new broth for Y. enterocolitica in order to improve high-throughput diagnostic PCR. This medium, named Yersinia-PCR-compatible-enrichment (YPCE) medium, was formulated on the following requisites: (i) the medium buffer system should mimic the buffer system in the PCR mixture, (ii) neither medium components nor metabolites produced by bacterial growth should be PCR-inhibitory, (iii) inclusion of selective components to restrict growth of competing background flora, and (iv) the medium should not interfere with fluorescence based detection used in real-time PCR. Using YPCE prior to conventional or real-time PCR allows detection of very low concentrations of Yersinia (<101 CFU/ml) in the presence of high levels of background flora. Finally, it is essential to optimize the DNA amplification conditions for a specific PCR sample matrix. A number of studies have proved that the choice of thermo-stable DNA polymerase and/or use of various amplification facilitators may dramatically improve the precision of diagnostic PCR by circumvent the presence of small amounts of PCR inhibitors and thus maintain an high amplification efficiency.112 The international standard EN ISO 20837:2006 specifies the requirements for sample preparation for qualitative detection of foodborne pathogens using PCR.114

36.2.3  Detection Procedures In this section we briefly comment on DNA-based methods for the detection of the three pathogenic Yersinia spp. and give detailed procedures for the detection of pathogenic Y. enterocolitica and Y. pseudotuberculosis by using real-time PCR based methods targeting the ail gene of both organisms. We recommend that DNA should be extracted using commercial purification kits when naturally contaminated samples are studied. The reason is that pathogenic Yersinia spp. are present in food in low numbers and considerable problems exist in getting the bacteria to grow in liquid media and thus only extraction procedures with high recovery rate will make detection possible.104 For pure cultures, boiling of bacteria in 100 µl of water for 10 min is enough. Also, commercial ready-to-use PCR master mixtures are available, both for use for conventional PCR and for realtime PCR. They can be recommended because they contain optimized concentrations of all components and are easy to handle. Y. pestis. For Y. pestis the detection assays targeting the pla gene appear to be the most sensitive, and the assay of

Molecular Detection of Foodborne Pathogens

Loiez and coworkers is a good candidate (Table 36.2). The real-time PCR assay is based on TaqMan technology coupled with automated DNA extraction and uses a minor groove binder-conjugated small DNA probe for DNA detection. Details of the method are given in the original publication.115 The only caveat in the assay is the location of the pla gene on the small pPst plasmid that can be lost during handling. Therefore, assays targeting also chromosomal Y. pestis-specific genes such as YP48, ypo2088, yihN, and entF3116–119 should be considered (Table 36.1). Y. pseudotuberculosis. The inv gene located in the chromosome and the virF/lcrF gene on the plasmid of Y. pseudotuberculosis have been used as targets for PCR (Table 36.2). Both genes can be detected using multiplex PCR with primers that amplify 295 bp inv and 591 bp virF/lcrF fragments according to Nakajima et al.120 Other specific target for detection of Y. pseudotuberculosis is the wzz-gene,55,118 however, in positive cases, the presence of Y. pestis has to be excluded, e.g., by one of the Y. pestis-specific targets (see above). Below we will, however, describe an ail gene-specific real-time PCR application for the detection of Y. pseudotuberculosis in food samples.121 Y. enterocolitica. A number of methods for specific detection of pathogenic Y. enterocolitica have been described (Table 36.4). The methods are designed to target either chromosomal or virulence plasmid-located genes or both; for detection using the chromosome as target molecule either ail, yst or 16S rDNA have been used and for target genes located on the virulence plasmid yadA, virF (lcrF) or yopN (lcrE). To be fully virulent pathogenic Y. enterocolitica strains must harbor a virulence plasmid. Since the plasmid may easily be lost, e.g., during subculture in the laboratory or storage over time, the virulence plasmid is not considered as a suitable target for the detection of the bacterium. In studies aiming to examine the presence of the virulence plasmid, it is therefore important to take into account the identified conditions (temperature, pH, calcium dependence etc.) that may retain the plasmid-bearing cells in the sample before PCR analysis.122 For that reason, we recommend the use of a method for detection of pathogenic Y. enterocolitica in food that targets the chromosomal located gene ail.123 The procedures. The same enrichment and DNA extraction procedure and PCR master mix can be used for detection of pathogenic Y. enterocolitica and Y. pseudotuberculosis. No reports are available on how to process food samples suspected to be contaminated by Y. pestis, however, a similar procedure as described below could also work for Y. pestis. The precautions when working with Y. pestis have been mentioned above (see Section 36.1.2.3). The overall detection scheme is illustrated in Figure 36.2. (i) Enrichment of food samples: (1) Dilute 25 g or 25 ml samples 1:10 in Tryptone soya broth (TSBY) medium (Tryptone soya broth 30  g/l supplemented with 0.6% yeast extract, pH 7.3). (2) Homogenize for 30 sec–1 min.

511

Yersinia

Day 0

25 g food + 225 ml TSBY Enrichment

Day 1

1 ml homogenate DNA extraction

25°C, 24 ± 3 hours

25 µl to CIN-agar

Real-time PCR Confirmation of PCR-positive results Day 2

Characterization of isolates

Figure 36.2  Overview of the proposed TaqMan-based real-time PCR method (in bold arrows). Steps indicated by dotted line arrows are suggested if verification of viable Yersinia bacteria is needed.



(3) Enrichment for 24±3 h at 25°C (4) Mix the enrichment, leave to sediment 15–30 min at 4–8°C. (5) Transfer 1 ml of homogenate for DNA extraction, see below. (ii) Confirmation of viable target bacteria: (1) Transfer 25 µl of the homogenate to a Cefsulodin Irgasan Novobiocin-agar (CIN) plate (prepared according to the instructions of the manufacturer). Spread the volume with a loop from side to side on the agar surface. Incubate at 30°C for 18–24 h. (2) If possible, pick from the CIN-agar plate presumptive Yersinia colonies or if the growth is confluent, draw a loop through a number of different colony types. Transfer the bacteria to one test tube containing 100 µl sterile distillated water before DNA extraction, see below. Note: One of the limitations of using PCR assays for detection of pathogens in food is that PCR will detect viable as well as nonviable organisms if the target is present in the sample. The inclusion of an enrichment step prior to PCR analysis will confine detection to viable culturable cells; in case of a positive PCR result, the presence of viable target bacteria can be confirmed by spreading a volume of the homogenate onto a CIN-agar plate and further proceeding as described above. If colonies can not be isolated, a positive PCR result can always be confirmed by reanalysis of the sample (using PCR). (iii) DNA extraction from homogenate or 100–200 µl of bacterial suspension: (1) Centrifuge 1 ml of homogenate or broth culture or 100–200 µl of bacterial suspension at 16,000×g for 10 min to sediment cells. Extract

DNA for example by using the DNeasy Blood and Tissue kit, mini-spin-columns (Qiagen Gmbh, Hilden, Germany). (2) Discard supernatant and resuspend pellet acco­r­ ding to the instruction of the manufacturer. Use 5 µl for PCR analysis. Store the rest for (possible) virulence plasmid investigation and/or for reanalysis. (3) If needed, DNA concentration can be measured. Note: Instead of using a “ready-to-use” kit, 100–200 µl of bacterial suspension may be treated as follows: add 10 µl of 0.8 M NaOH to 100 µl of bacterial suspension and heat 10 min at 70–75°C. Then, add 24 µl of 1:1 mixed solutions of 1 M Tris and 0.8 M HCl. Mix and use 5 µl for PCR analysis. Store the rest for (possible) virulence plasmid investigation and/or for reanalysis. (iv) Real-time PCR: The pipetting scheme, real-time PCR conditions and primer sequences based on the ail-genes of Y. enterocolitica123 and Y. pseudotuberculosis are given in Tables 36.5 and 36.6.

36.2.4 Quality Assurance Aspects When Using PCR The increased interest in implementing quality assurance and control measures in laboratories using PCR will ensure successful and reliable PCR results. According to ISO standard 22174:2005, inclusion of controls to monitor the analysis performance is considered obligatory and must be added in each PCR run when analysing for pathogens in food samples.124 A number of controls exist, for example, negative controls and amplification controls. A negative PCR control is used to detect any PCR-fragment carry-over contamination. In a negative PCR control the test sample volume is replaced by sterile distilled water. An amplification control serves to control the amplification and will monitor for falsenegative PCR results. Thus, a negative PCR result obtained both by the amplification control and the target bacteria is an indication of a PCR inhibition. These samples can be rerun after dilution of the sample by 1:10 (or more). The concentration of the added control DNA fragment should be as low as possible in order to detect even small inhibitions and at the same time give a statistical reproducible positive result. Both internal (IAC) and external amplification controls exist. Furthermore, according to ISO 20838:2006125 confirmation of the identity of a PCR product obtained by conventional PCR should be undertaken by an appropriate method other than size determination, for example by DNA sequencing, by hybridization with specific DNA probes or by carrying out restriction analysis of the PCR product. Confirmation of positive PCR result by culture and characterizing the isolates by conventional methods (i.e., biochemical tests, serotyping, etc.) is encouraged, but usually difficult to achieve due to the low detection efficiency of culturing methods.

512

Molecular Detection of Foodborne Pathogens

Table 36.5 Pipetting Scheme for TaqMan Real-Time PCR Reagents and Real-Time PCR conditions for the Detection of ail-Gene Component (Stock Conc.)

Volume/Reaction (Final Conc.)

2.5 µl (1×) 10× TaqMan buffer A MgCl2 (25 mM)a 3.5 µl (3.5 mM) dNTPs (50 mM)a 0.1 µl (200 µM) 0.25 µl (0.02 U/µl) AmpliTaqGold® (5 U/µl)a Primer 1 (10 µM) 0.75 µl (300 nM) Primer 2 (10 µM) 0.75 µl (300 nM) Probe (20 µM) 0.25 µl (200 nM) Approx. 100 copies IACb Test sample 5 µl Sterile distilled water Add 25 µl Real-time PCR cycling parameters (optimized for the 7500 ABI system) One cycle of 95°C for 10 min 45 cycles of 95°C for 15 sec and 60°C for 1 min a

a

b

Comments Applied Biosystems, Foster City, CA

For sequence see Table 36.6 For sequence see Table 36.6 For sequence see Table 36.6

These components can be replaced by the 10× TaqMan® Universal PCR Master Mix (Applied Biosystems, Foster City, CA) or by similar buffers from other suppliers. When naturally contaminated samples are tested it is strongly recommended to use the master mix with the inclusion of dUTP instead of dTTP. (The final concentration of the Master mix components: 1× TaqMan Universal PCR mix, 900 nM of each primer, 200 nM of probe. Test sample, 5 µl. Total volume, 25 µl). Two internal amplification controls (here named IPC and IAC) are presented here: (i) A commercially available TaqMan® Exogenous Internal Positive Control (Applied Biosystems, Foster City, CA). The reagent kit includes primers, a Vic™probe, IPC target DNA and blocking solution. The IPC target DNA needs to be diluted ten times to achieve a copy number of approximately 100 per PCR reaction. The PCR product length is not given to the customer. (ii) An open formula pUC19-based internal amplification control IAC developed by Fricker et al.184 can be used. Approximately 50–100 copies of target DNA (pUC19) are used per PCR reaction. The size of this IAC is 119 bp.

Table 36.6 Primers and Probes used for Real-Time PCR Detection of Pathogenic Y. enterocolitica and Y. pseudotuberculosis Primer or Probe Y. enterocolitica F-real 10A R-real 9A Ye probed Y. pseudotuberculosis F-Yps 1 R-Yps 2 Probed

Sequence (5´–3´)

Positiona

Melting Temperature (ºC)b

atgataactggggagtaataggttcg cccagtaatccataaaggctaacatat FAM-tctatggcagtaataagtttggtcacggtg atct-TAMRAc (Note: These primers will also detect Y. Pestis CGTCTGTTAATGTGTATGCCGAAG GAACCTATCACTCCCCAGTCATTATT VIC-CGTGTCAAGGACGATGGGTACAAGTTGGTAMRAc

2379–2404 2516–2541 2436–2469

63.0 61.5 68.1

163 bp

1395870-1395893 1396001-1396026 1395929-1395956

59.6 58.8 69.5

157 bp

Amplicon Size

Nucleotide positions correspond to the Y. enterocolitica O:3 ail-gene sequence (GenBank accession no AJ605740) and to Y. pseudotuberculosis YPIII complete genome sequence (GenBank accession no CP000950.1). b Calculated with the Primer Express program, version 3.0 (Applied Biosystems, Foster City, CA). c TAMRA™; FAM™ ; VIC® (Applied Biosystems, Foster City, CA). d The Y. enterocolitica probe can be exchanged for the MGB probe: FAM-TGACCAAACTTATTACTGCCATA-MGB. The Y. pseudotuberculosis probe can be exchanged for the MGB-probe: NED-ATGCTCAAAGTCGTGTCAA-MGB. MGB, minor groove binder. (Probes were purchased at Applied Biosystems, Foster City, CA) Source: Lambertz, S.T., Nilsson, C. and Hallanvuo, S. Appl. Environ. Microb. 74, 6465, 2008; Lambertz, S.T. et al., Appl. Environ. Microb. 74, 6060, 2008. a

513

Yersinia

36.3 Conclusions and Future Perspectives This chapter highlights some of the advantages and disadvantages with the current molecular methods available for detection of pathogenic Yersinia spp., and in particular PCRbased methods. Quantification of pathogens in naturally contaminated samples is still complicated and further research is needed to establish the PCR methodology that will ensure detection of live cells only. In the future development of diagnostic PCR, research in sample preparation is likely to expand in response to the growing demand for rapid, robust and simple PCR protocols. A future challenge for sample preparation is to design PCR protocols that integrate DNA extraction and amplification in an automated manner. Sample preparation influences both the specificity and the sensitivity of the overall PCR protocol. To minimize these variations it is necessary to thoroughly optimize the pre-PCR processing conditions. From a quality point of view as few steps as possible should be included in the process of diagnostic PCR. Using unconventional DNA polymerases and amplification facilitators in combination with optimized sampling conditions, reduces the amount of manual handling involved in the analysis of pathogenic bacteria. On the other hand, as the number of available (bacterial) genome sequences is increasing the possibility to define novel biomarkers for PCR and microarray assays will definitely increase in the future. This, along with the development of novel detection devices will most likely allow the design of high-throughput systems that can cost-efficiently detect simultaneously all major pathogens from different types of samples. Real-time PCR is a powerful advancement of the basic PCR technique. In contrast to conventional PCR, real-time PCR facilitates automation, computerisation and quantification of nucleic acids. A significant improvement introduced by real-time PCR is the increased throughput and reduced carry-over contamination. This is largely due to reduced cycle times, removal of separate post-PCR detection procedures and use of sensitive fluorescence detection equipment, allowing earlier amplicon detection. Future improvements in real-time PCR assays that focus on increasing the automation of the entire process will enhance the usefulness of this method in laboratory diagnostic, epidemiological studies, and the food industry. In addition high-throughput technologies such as DNAmicroarray, mass spectrometry-PCR and whole genome sequencing platforms will strengthen the portfolio of molecular based diagnostic tools for pathogenic Yersinia spp. Those methods will allow rapid molecular characterization and also the possibility to determine antibiotic resistance of the isolate. Due to the many on-going bio preparedness programs, these high-throughput-based technologies have been applied for the detection and characterization of Y. pestis. Several Y. pestis genomes, as described in Figure 36.1A, have already been completed funded by bio preparedness research

programs. However all these methods require optimized sampling and sample preparation prior to analysis. A future challenge of available high-throughput Yersinia detection methods, including real-time PCR, is to develop integrated workflows adapted to food safety concerns.

Acknowledgments The work of MS has been supported by the Academy of Finland (projects 114075 and 104361) and the European Union Network of Excellence “EuroPathoGenomics” (contract LSHB-CT-2005-512061). The work of PR has been supported by the Swedish Research Council for Enviroment, Agricultural Sciences, and Spatial Planning (Formas).

References





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Molecular Detection of Foodborne Pathogens 180. Ikeda, M. et al., Rapid and simple detection of food poisoning bacteria by bead assay with a microfluidic chip-based system, J. Microbiol. Methods, 67, 241, 2006. 181. Magliulo, M. et al., A rapid multiplexed chemiluminescent immunoassay for the detection of Escherichia coli O157:H7, Yersinia enterocolitica, Salmonella typhimurium, and Listeria monocytogenes pathogen bacteria, J. Agric. Food Chem., 55, 4933, 2007. 182. Chaney, R., Rider, J. and Pamphilon, D., Direct detection of bacteria in cellular blood products using bacterial ribosomal RNA-directed probes coupled to electrochemiluminescence, Transfus. Med., 9, 177, 1999. 183. Cloak, O.M. et al., Isolation and detection of Listeria spp, Salmonella spp and Yersinia spp using a simultaneous enrichment step followed by a surface adhesion immunofluorescent technique, J. Microbiol. Methods, 39, 33, 1999. 184. Fricker, M. et al., Diagnostic real-time PCR assays for the detection of emetic Bacillus cereus strains in foods and recent food-borne outbreaks, Appl. Environ. Microbiol., 73, 1892, 2007.

Section IV Foodborne Fungi

37 Alternaria

Dongyou Liu, Stephen B. Pruett, and Cody Coyne Mississippi State University

Contents 37.1. Introduction..................................................................................................................................................................... 521 37.1.1. Classification, Morphology, and Biology.......................................................................................................... 521 37.1.2. Clinical Presentation, Pathogenesis, and Epidemiology................................................................................... 522 37.1.3. Laboratory Diagnosis........................................................................................................................................ 523 37.2. Methods........................................................................................................................................................................... 524 37.2.1. Sample Preparation............................................................................................................................................ 524 37.2.2. Detection Procedures........................................................................................................................................ 524 37.2.2.1. PCR and Sequencing Analysis of ITS-5.8S Gene Region................................................................ 524 37.2.2.2. PCR Detection and Quantification of AM-toxin Gene.................................................................... 525 37.3. Conclusions and Future Perspectives.............................................................................................................................. 525 References.................................................................................................................................................................................. 526

37.1  Introduction 37.1.1 Classification, Morphology, and Biology The genus Alternaria (Nees) Wiltshire was established on the basis of the conidial morphology in 1933 to cover a group of filamentous fungi in the phylum Ascomycota, the family Dematiaceaee, that had previously been known as Alternaria tenuis and Torula alternate.1 Morphologically, members of the genus Alternaria form grey to blackish-brown or black colonies. The colonies of the species A. alterniria (syn. A. tenuis) are filamentous, grey, dark brown or black, fast growing, black to olivaceous-black or greyish, and suede-like to floccose. Alternaria conidiophores (measuring 2–6 nm × 20–50 nm) are single or in small groups, straight or curved, sometimes geniculate, showing scars after they are detached. Microscopically, they are characteristically simple, sometimes branched, short or elongate conidiophores with branched acropetal chains (blastocatenate) of multicelled conidia (dictyoconidia). Conidia (or spores, measuring 9–18 nm × 20–63 nm) are ellipsoidal, ovoid, obclavate, obpiriform, or club-shaped, often with a short conical or cylindrical beak (measuring 2–5 nm in diameter), which is pale brown, smooth-walled or verrucose, tapering to an apex or blunt, about one third or one quarter of the conidial length. The surfaces of conidia are usually smooth, but some display small rounded warts presenting a slightly rough surface. Alternaria conidia (spores) with one to eight transverse septa and and to two longitudinal or oblique septa in each cell are golden-brown to brown in color with a beak that is pale in appearance. Conidia are formed by blastic ontogeny as outgrowths of protoplasm through a defined apical pore in the conidiogenous cell.2 One characteristic feature useful for discrimination of Alternaria species is the presence or absence of chains and the

number of conidia in these chains.2 Alternaria species having ten conidia (spores) or more are known as Longicatenatae; those with three to five conidia (spores) as Brevictenatae; and those with single conidia (spores) as Noncatenatae. Another diagnostic feature for Alternaria species is the formation of an apical beak of varying length in the conidia and the nature of the transition from body to beak and septation. When conidia proliferate as secondary conidiophores to originate new conidial chains, a pseudorostrum may be present. Alternaria may be confused with genera Stemphylium and Ulocladium because they possess a similar type of spore (porospores). However, Stemphylium is distinguished from Alternaria by its percurrent prolifereation, and Ulocladium by the narrow base of conidia which differs from the broad base in Alternaria and by the lack of a beak-like configuration at the apex of conidia. In addition, Stemphylium, and Ulocladium do not produce conidia in chains (Stemphylium) or they only have very short chains (Ulocladium).3 Alternaria spp. are dematiaceous molds that are cosmopolitan and diverse, with the number of Alternaria species being estimated to be several hundred, and approximately 80 species being described so far. Nevertheless, due to their highly varied morphological features, it is possible that several Alternaria that have been described as distinct species may in fact actually belong to a single identical species. Some species of Alternaria are the asexual anamorph of the ascomycete Pleospora while others are speculated to be anamorphs of Leptosphaeria. Alternaria spp. are plurivorous on soil, fibers, food, and vegetable hosts (tress, shrubs, and crops). Many Alternaria species grow saprophytically in a range of habitats (deriving energy from their cellulytic activity), are ubiquitous agents of decay and are pathogenic to vegetable host and stored grains, seeds, and fruits. However, there are 521

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eight Alternaria species (i.e., A alternata, A brassicicola, A chartarum, A stemphylioides, A dianthicola, A infectoria, A pluriseptata, and A tenuissima) that have been implicated as human pathogens. Additionally, A. iridis is mentioned as a strain responsible for creating allergic reactions. As an epiphyte, Alternaria is capable of penetrating to the level of host subepidermal tissues asymptomatically and colonizing substomatal chamber, where it develops into single five to seven celled fusiform hyphae that assimulate nutrients from the host and derive protection from desiccation and mycophagous invertibrates. After harvest of fresh foods, viability of Alternaria may persist and cause deterioration of fruits and vegetables. Having the ability to sustain low humidity levels, which inhibit the growth of Aspergillus and Penicillium species, Alternaria thrives on stored cereal grains. However, Alternaria generally does not tolerate 0.15 or higher humidity levels. Besides being mycotoxin producers, Alternaria species secrete a large number of enzymes including cellulose, pectin methylgalacturonase, pectin methylase, pectin methylesterase, and cutinases, some of which have been utilized as industrial raw materials and pharmaceutical reagents. The optimal growth temperature for Alternaria is 25–28°C; and the maximal growth temperature is 31–32°C. When incubated at 25°C (77°F), Alternaria replicates quickly and forms a grayish-white colony that within 5 days becomes greenish-black, with pigmented olivaceous-brown (dematiaceous) hyphae, conidiophores, and conidia. The most suitable growth media for Alternaria are potato dextrose agar (PDA), cornmeal agar or malt extract agar (MEA). As Alternaria species easily lose their ability to sporulate in culture, it is recommended to incubate Alternaria under near ultra-violet in order to maintain sporulation activity.

37.1.2 Clinical Presentation, Pathogenesis, and Epidemiology Clinical presentation. While many Alternaria species are important plant pathogens that cause surface discoloration initially followed by disruption of plant internal structures, reducing their yield, viability, quality, nutritional, and economic values,4 a small number (at least eight species) are capable of inducing human health disorders (called alternariosis or alternariatoxicosis, which may include superficial, cutaneous, subcutaneous, corneal or systemic) in humans and animals.5–9 Firstly, some Alternaria species are opportunistic pathogens that grow on skin and mucous membranes including on the eyeballs (ocular mucosa) and within the respiratory tract, oral, and sinus cavities, causing onychomycosis, ulcerated cutaneous infections, mycotic keratitis, respiratory diseases, especially in those with underlying diseases/conditions that compromise the host’s immune function (e.g., AIDS).10–20 The early clinical sign of Alternaria-induced cutaneous ulcerations is a macula, which becomes a squamous, crustlike, reddish-violaceous, ulcerated nodule. Biopsies of these lesions reveal a well-delimited dermal granuloma, while culture analysis confirms the presence of the fungi. Secondly, Alternaria spores can elicit a hypersensitivity

Molecular Detection of Foodborne Pathogens

pneumonitis (woodworker’s lung disease), chronic rhinosinusitis (or paranasal sinusitis) and an immediate-type hypersensitivity-type 1 (IgE-mediated) extrinic asthma (hay fever). Acute symptoms include edema and bronchiospasms, while chronic cases may develop pulmonary emphysema. Thirdly, Alternaria can be a causative agent for osteomyelitis, peritonitis, cellulitis, and other visceral infections in patients following organ transplantation21 and those managed by continuous ambulatory peritoneal dialysis. Fourthly, Alternaria species are among the causative agents that can precipitate otitis media in agricultural field workers. Pathogenesis. Alternaria produces dark, melanin (i.e., dihydroxynaphthalene melanin)-rich mycelium and conidia that are resistant to radiation and adverse environmental conditions. Being easily dispersed by wind, Alternaria spores colonize plants via wounds to the pericarp layer. About 7–10 days after host colonization, Alternaria sporulates and generates conidia for several weeks. The undesirable effects of Alternaria species on plants include the promotion of spots or blights in leaves; spots and cankers in stems and twigs; damping off in seedlings; fruit, tuber, root, and seed rots.22 Alternaria spores may remain in the harvested fruits and grains, and geminate in propitious temperature and humidity conditions in storage. Besides causing at least 20% of agricultural spoilage and diminishing the value of the commodities, the geminating spores of Alternaria species secrete both host-specific and general phytotoxins/mycotoxins and secondary metabolites, which amount to over 70 in total, contributing to their pathogenicity on plants and toxicity to humans and animals as food and feed contaminants.23–28 The major mycotoxin produced by Alternaria species come under three distinct structural groups: (i) dibenzopyrone derivatives including alternariol (AOH), alternariol monomethyl ether (AME), and alternuene (ALT); (ii) perylene derivatives (alteroxins ATX-I, II, and III); and (iii) tetramic acid derivative, tenuazonic acid (AT).26 Tenuazonic acid (AT) from A. alternata and other toxic metabolites are associated with hypersensitivity pneumonitis, sinusitis, deratomycosis, onychomycosis, subcutaneous phaeohyphomycosis, and invasive infections. They also cause extrinsic asthma (immediate-type hypersensitivity: type I) characterized by acute symptoms of edema and bronchiospasms, in addition to chronic symptoms of pulmonary emphysema. Some other mycotoxins and secondary metabolites show mutagenic and teratagenic properties, and are linked to certain forms of cancer. Being one of the most common and potent indoor and outdoor airborne allergens, Alternaria spores (having sizes between 20 and 200 μm in length and 7–18 μm in width) can be easily deposited within the nose, mouth, and upper respiratory tract, causing childhood asthma and baker’s asthma. As a complex identity comprising several heterogenous species, A. alternata is more commonly isolated from human infections than other Alternaria spp., which include A. alternata, A. brassicicola, A. chartarum, A. stemphylioides, A. dianthicola, A. infectoria, A. pluriseptata, and A. tenuissima. Epidemiology. Alternaria species are widely distributed in the environment, and are present in soil, wood, and

Alternaria

decomposing plant debris, especially in tropical and subtropical climates. Alternaria species produce abundant airborne spores and mycelium debris during spring, summer, and autumn due to the degradation of leaves and other vegetations. In indoor plants, Alternaria species produce spores all year around. Exposure to airborne spores and mycelium debris of the Alternaria species (through inhalation and traumatic implantation) leads to invasive sinusitis and allergic fungal sinusitis in immunocompetent individuals,29 and systemic phaeohyphomycosis (in the form of cerebral infection) and other localized “deep” forms of the disease (e.g., arthritis and endocarditis) in immunocompromised individuals.

37.1.3 Laboratory Diagnosis Alternaria species such as A. alternata, A. infectoria, A. tenuissima, and A. chartarum are often localized on skin as a result of direct, traumatic inoculation, and induce difficultto-recognize lesions that vary in size and appearance since they can present as crusted lesions erythematous macules or subcutaneous nodules. Considering that several other fungal infections such as blastomycosis, sporotrichosis, cryptococcosis, and other subcutaneous mycoses may present with similar clinical signs, and that Alternaria infection may be misdiagnosed histopathologically as a yeast infection,31 it is necessary to use laboratory techniques for the diagnosis of Alternaria infections. Conventional techniques. Traditional methods for Alternaria identification are dependent on examination of the morphological characteristics of its colony, vegetative hyphae, conidia, conidiophores, the spatial patterns of sporulation and host or substrate. Due to their inherently rapid growth characteristics, Alternaria colonies reach a diameter of 3–9 cm in size after 7 days on potato glucose agar at 25°C. Colonies appear flat, downy to woolly, and are covered with grayish, short, aerial hyphae. Initially their color is grayish white with the surface subsequently turning greenish black or olive brown with a light border at a later point in time while the reverse side reveals brown to black pigment production. Microscopically, Alternaria hyphae and conidiophores appear septated and brown in color, with simple or branched large conidia (7–10 × 23–34 µm) possessing both transverse and longitudinal septations. The conidia are obclavate or muriform in shape with club-like appearance and may appear smooth or have a rough looking texture. The use of hematoxylin and eosin (H&E) stain enables visualization of dark-colored filamentous hyphae in sections of infected tissue (especially cutaneous mycoses); and Fontana-Masson silver stain may be employed to specifically detect melanin if dark-colored hyphae are not obvious. Specimens of subcutaneous mycoses may show large, hyaline, yeast-like cells. Occasionally, A. alternata and A. infectoria produce nonpigmented hyphae which may lead to misidentification.32 Alternaria species secrete over 70 mycotoxins, that are harmful or carcinogenic to plants and animals which are responsible for causing considerable economic losses to growers and the food-processing industry. High-performance

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liquid chromatography (HPLC), enzyme-linked immunosorbent assay (ELISA) and other phenotypic techniques have been successfully applied to detect Alternaria toxins.33,34 The glycoprotein, ALT a 1 is the major allergen produced by A. alternate that has at least five isoelectric variants between pI 4.0 and 4.3 with apparent molecular masses between 25-kDa and 50-kDa. ALT a 1 is responsible for inducing 70% of IgE-binding activity in sera from human patients with A. alternate exposure. By using monoclonal and polyclonal antibodies directed against Alt a 1, an ELISA has been developed that is capable of reproducible and sensitive detection of A. alternata, with a detection limit down to 0.5 ng/mL and a practical detection range of 0.5–50 ng/mL. Excellent correlation between Alt a 1 content of 13 A. ­alternata extracts and their IgE-binding activity was observed. This sensitive and specific ELISA for Alt a 1 allows the quantification of A. alternata and provides a reliable indication of the allergenic activity of the whole extract.34 Molecular techniques. While analysis of morphological, biochemical, and serological features permits accurate identification of Alternaria spp., the whole process is technically demanding, time-consuming and variable accurate due to subtle morphological differences within the species. Indeed, a week or more is often required for in vitro cultivation and appropriate sporulation medium is needed to obtain optimal conidiation.35,36 To facilitate early diagnosis and effective treatment of alternariasis, molecular techniques have been increasingly applied as a rapid, sensitive, and specific alternative to phenotype-based procedures in the detection and identification of Alternaria species.37,38 Similar to other eukaryotes, Alternaria ribosomal RNA (rRNA) genes consist of three subunits (i.e., 18S, 5.8S, and 28SS). Between the 18S and 5.8S and between the 5.8S and 28S ribosomal DNA (rDNA) gene subunits are intergenic transcribed spacer regions (ITS1 and ITS2) that are not translated into rRNA. While rRNA genes are highly conserved, the ITS regions show sequence divergence, with sufficient internal variations to be exploited for laboratory diagnostic purposes relevant to Alternaria infections. By using PCR primers targeting the ITS1 and ITS2 regions of Alternaria 5.8S rDNA gene, rapid, and simultaneous testing of multiple samples can be achieved at a low cost.33,39–46 Phylogenetic analysis of the sequence data by the Neighbor-joining method showed that the seven toxin-producing fungi belong to a monophyletic group together with A. alternata. By contrast, A. dianthi, A. panax, A. dauci, A. bataticola, A. porri, A. sesami and A. solani are morphologically distinct A. alternata, and they can be clearly separated from A. alternata by phylogenetic analysis of ITS variations. Exploiting the difference in the ITS domains between the two most important, longicatenate species, A. alternata and A. infectoria (which is one of the most common clinical Alternaria species that has a low degree of melanization and contains a 26-bp insert in its ITS1), de Hoog and Horre3 have devised a PCR-restriction fragment length polymorphism (RFLP) assay that can distinguish between these two species. Although several taxa inhabiting plants such as A. longipes

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on tobacco, A. mali on apples, and common saprobic species A. tenuissima, cannot reliably be distinguished from A. alternata using this method, these species rarely cause human disease and are not found in clinical samples. Dubois et al.45 described a case of cutaneous alternariosis that created papular, crusted lesions on the left elbow and the right knee of a 74-year-old man treated with corticotherapy for myasthenia. A dematiaceous fungus was isolated and identified as a member of the A. infectoria species-group by PCR amplification in concert with sequencing of the ITS gene encoding nuclear ribosomal DNA, and the mitochondrial small subunit ribosomal DNA domain. Laich et al. 47 also reported a PCRRFLP assay targeting the ITS domain of 5.8S and 26S regions of nuclear rDNA for the detection of A. alternata in corneal scrapings collected from keratitis lesions. Additionally, A. alternata isolates can be characterized using random amplified polymorphism of DNA (RAPD).48,49 Since Alternaria species are capable of producing a large array of mycotoxins, it is useful for the food industry to know if toxigenic fungi are present in raw vegetables and animal products. Thus it is essential to have rapid and accurate ­methods available for detection and identification of Alternaria organisms that are pathogenic to plants as well as the production of toxic metabolites. Such an achievement would facilitate the prompt implementation of appropriate corrective actions as needed. By using an AM-toxin I primer set, it has been shown that Alternaria isolates producing the plant pathogenic ­tentoxin and AM toxin I can be detected with great sensitivity (e.g., 4 pg of DNA by real-time PCR).50,51

Molecular Detection of Foodborne Pathogens

grinding in liquid nitrogen, mechanically disrupted at ambient temperature, sonicated, subjected to glass bead milling, or serially freezing and thawing. Haugland et al.40,54 found that glass bead milling is more effective than other methods for recovery of PCR templates from Alternaria conidia. If conventional DNA isolation approach is utilized, the disrupted/ homogenized sample is resuspended in extraction buffer (50 mM Tris-HCl, 50 mM ethylenediaminetetraacetic acid, 3% sodium dodecyl sulfate, 1% 2-mercaptoethanol), and extracted with phenol-chloroform-isoamyl alcohol (25:24:1). The DNA remaining in the supernatant is then precipitated with isopropanol and washed with 70% ethanol. The final DNA pellet is dissolved in 10 mM Tris-HCl pH 7.5 and the concentration measured as a function of UV absorbance at 260 nm. Alternatively, commercial kits such as Qiagen DNeasy® Plant Minikit (Qiagen) or Fast DNA Spin Kit for Soil (BIO101, Carlsbad, CA) can be utilized to purify Alternaria DNA in the absence of biohazard chemicals.

37.2.2  Detection Procedures

37.2  Methods

37.2.2.1 PCR and Sequencing Analysis of ITS-5.8S Gene Region Principle: Ferrer et al.41 employed a pair of universal primers from the ITS-5.8S region of fungal DNA (ITS1: 5′ TCC GTA GGT GAA CCT GCG G 3′ and ITS4 (5′ TCC TCC GCT TAT TGA TAT GC 3′)39 to achieve PCR amplification of a 600-bp fragment. After elucidation of the nucleotide sequence of this fragment and comparison with sequences in the GenBank database using the BLAST alignment program, Alternaria species could be precisely determined.

37.2.1 Sample Preparation

Procedure:

Sample processing. Food samples such as grains and apples suspected of being infected by Alternaria can be used directly for DNA extraction after surface sterilization (e.g., soaking in 0.4% hypochlorite solution for 2 min and serially washed three times with sterilized distilled water for 1 min), although it can be helpful to obtain in vitro isolates of Alternaria organisms for further phenotypic and molecular confirmation.52,53 Human clinical samples (e.g., skin biopsies) often need to go through in vitro culture process prior to molecular detection. The in vitro culture can be done on Sabouraud’s dextrose agar, potato dextrose, and/or MEA at 30 and 35°C for 7 days, where upon yellowish-white Alternaria colonies develop that have an olive-green undersurface emerge. Upon microscopic examination of lactophenol cotton blue-stained smears, the fungus often shows melanized septate hyphae. Subcultures on Sabouraud dextrose-chloramphenicol agar, MEA and/or and water agar at 25°C for 6 days may be performed to enhance sporulation to improve detection of obclavate or pyriform conidia that contain transverse and longitudinal or oblique septa. DNA isolation. Alternaria DNA can be extracted from conidia or food samples using conventional procedures or commercial kits. Conidia or food samples may be homogenized by

(1) Prepare PCR mixture (50 µl) containing 6 µl of 25 mM MgCl2, 5 µl of PCR buffer without MgCl2, 200 µM each deoxynucleoside triphosphate (dNTPs), 25 pmol of each primer (ITS1 and ITS4), 1 U of Taq DNA polymerase, and 2 ng of DNA template. (2) Conduct PCR amplification for one cycle of 95°C for 5 min, 35 cycles of 95°C for 30 sec, 55°C for 1 min, and 72°C for 1 min; and one cycle of 72°C for 6 min. (3) Run 10 µl of each amplified product on a 2% agarose gel in 1 × Tris-borate-EDTA buffer. Include molecular weight ladders on one of the lane (pBR322 DNA/ BsuRI or Gene Ruler 100-bp DNA Ladder Plus). Visualize the PCR band with ethidium bromide stain under UV illumination. (4) Excise the PCR band from the gel, purify using the GeneClean II kit (Bio101) and sequence in both directions (with the same primers) using the BigDye terminators Ready Reaction Kit (PE Applied Biosystems, Foster City, CA) on an ABI Prism automated DNA sequencer (model 377, version 2.1.1; Applied Biosystems). Align the resulting nucleotide sequence with those in the GenBank database using the BLAST program.

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Alternaria

Note: PCR amplification of the ITS-5.8S rDNA gene using specific fungal primers ITS1 and ITS4 results in the formation of a 600-bp fragment. The homology of the ITS5.8S rDNA sequences among strains of the same species approaches 99.5%. 37.2.2.2 PCR Detection and Quantification of AM-toxin Gene Principle: Andersen et al.51 described the use of primers targeting the 3′ end of the AM-toxin synthase gene (i.e., AM-forward: 5′-ATATCGTTTTCGGCCGCAC pos 13343– 13361 and AM-reverse: 5′-AACGGGTCCATGTAACCTAG pos 13739–13758) for standard PCR detection and realtime PCR quantification of apple pathogenic and toxigenic Alternaria. The method enabled detection of 4 pg of DNA from apple material based on real-time format. Procedure: (i) Standard PCR detection: (1) Prepare PCR mixture (50 μl) consisting of 3 mM MgCl2, 200 μM dNTPs, 2.5 U Taq polymerase, 1 × PCR buffer, 0.5 μM AM-forward and AM-reverse primer, and 2 ng genomic DNA. (2) Conduct PCR amplification on a thermocycler for one cycle of 94°C for 2 min; 40 cycles of 94°C for 30  sec, 53°C for 1 min and 72°C 1 min; and one cycle of 72°C for 10 min. (3) Digest 10 μl of the PCR product with 10 U of restriction enzyme HincII in 1 × buffer, 0.25% bovine serum albumin at 37°C for 2 h. (4) Separate the digested product on 1.5% agarose gel containing 0.5 μg/ml ethidium bromide, and visualize with UV illuminator. Note: A specific PCR fragment of 436 bp is obtained from Alternaria DNA using AM-forward and AM-reverse primers. As restriction enzyme HincII cuts at position 13429 in the AM-toxin gene and at position 351 in the resulting PCR fragment, an expected DNA fragment of 351 bp (and a fragment of 85 bp, which is not detectable in the gel) confirms the presence of the AM-toxin gene. Out of 20 Alternaria strains examined, eight were positive for the AM-toxin gene.51 (ii) Real time PCR quantitation: (1) Prepare real time PCR mixture containing 4 mM MgCl2, 0.5 μM AM-forward and reverse primer, 2 μl Light cycler DNA master SYBR Green I, 1 × DNA buffer, 1 × DNA polymerase master mix, and 2 ng of genomic DNA. (2) Conduct PCR amplification using a Light cycler™ real-time PCR apparatus (Roche Diagnostics) with the following conditions: one cycle of 95°C for 2 min, 40 cycles fo 95°C for 5 sec, 63°C for 5 sec, and 72°C for 12 sec; and one cycle of 72°C for 10 min.

(3) Plot the crossing point (i.e., the point at which the PCR product is detectable by the laser system of the equipment) against the amount of DNA added to each sample. Note: The detection limit of this real-time PCR was shown to be at least 4 pg of fungal DNA corresponding to approximately 75 genomes using an estimated Alternaria genome size of 30 Mb.55

37.3 Conclusions and Future Perspectives The genus Alternaria encompasses a diverse group of filamentous fungi that are distributed ubiquitously within many environments. Morphologically, Alternaria species differ from other dematiaceous fungi by possessing chains of pigmented conidia that often show transverse and longitudinal septations and a distinct apical beak.1,2 Besides leading a saprophytic existence on decaying vegetations, Alternaria species are capable of invading and colonizing plants through spores that are easily dispersed by wind. Within the plants, Alternaria sporulates and produces conidia that cause discoloration in leaves, fruit, seed, and roots (what are seed rots...???), reducing the nutritional values of agriculture products. In addition, Alternaria species generate over 70 mycotoxins and secondary metabolites that contaminate food products and cause foodborne illnesses in human and animals. Of the 80 or so species that have been described, at least eight Alternaria species (i.e., A alternata, A brassicicola, A chartarum, A stemphylioides, A dianthicola, A infectoria, A pluriseptata, and A tenuissima) have been shown to cause superficial, cutaneous, subcutaneous, corneal or systemic infections in humans, particularly those individuals with suppressed immune function or underlying diseases. Further, Alternaria spores are allergens that often induce asthma (hay fever), rhinitis and other disorders of immunity in susceptible population groups. In order to design and implement suitable treatment strategies against alternariosis, and prevent foodborne diseases due to Alternaria mycotoxins, it is vital that Alternaria organisms can be accurately identified. Although conventional techniques based on in vitro culture and microscopic examinations can provide a definitive diagnosis, they suffer from the shortcomings of being slow and sometimes variably accurate in addition to requiring specialized laboratory skills. The application of nucleic acid-based techniques such as PCR procedures makes rapid, sensitive, and specific identification and detection Alternaria species feasible. To date, most molecular assays have targeted Alternaria rRNA and ITS1 and ITS2. Because of the relative genetic conservation in these regions, the resulting PCR products are often of similar size, which requires extra steps such as DNA sequencing and restriction enzyme digest to distinguish among related species. Therefore, future characterization of genes unique to individual Alternaria species will facilitate development of species-specific tests that negate the need to perform additional time-consuming steps after PCR amplification.

526

Given that Alternaria species produce a large number of mycotoxins that are toxic and carcinogenetic to humans and animals, it is critical to utilze rapid and sensitive techniques for detecting toxins and their corresponding genetic sequences. In effect, this will help reduce potential foodborne illnesses resulting from the consumption of food products containing Alternaria and its mycotoxins. Use of PCR assays targeting AM-toxin synthase offers a valuable means for identification of toxigenic Alternaria.51 Further development of molecular assays targeting other toxin genes in Alternaria species will help ensure that the food products we consume are free of Alternaria mycotoxin contamination.

References









1. Simmons, E.G. Alternaria taxonomy: current status, viewpoint, challenge. In: Alternaria: Biology, Plant Diseases and Metabolites, pp. 1–36. Chelkowski, J. and Visconti, A. (eds.). Elsevier, Amsterdam, The Netherlands, 1992. 2. Lopez, S.E. and Cabrel, D. Alternaria. In: Encyclopedia of Food Microbiology, pp. 42–49. Robinson, R.K., Batt, C.A. and Patel, P.D. (eds.). Academic Press, San Diego, CA, 2000. 3. de Hoog, G.S. and Horre, R. Molecular taxonomy of the Alternaria and Ulocladium species from humans and their identification in the routine laboratory. Mycoses, 45, 259, 2002. 4. Serdani, M. et al. Characterisation of Alternaria species-groups associated with core rot of apples in South Africa. Mycol. Res., 106, 561, 2002. 5. Gerdsen, R. et al. Sporotrichoid phaeohyphomycosis due to Alternaria infectoria. Br. J. Dermatol., 145, 484, 2001. 6. Verma, K. et al. Post-LASIK infectious crystalline keratopathy caused by Alternaria. Cornea, 24, 1018, 2005. 7. Akman et al. Cutaneous alternariosis in a patient with systemic lupus erythematosus. Lupus, 16, 993, 2007. 8. Bonatti, H. et al. Alternaria alternata soft tissue infection in a forearm transplant recipient. Surg. Infect. (Larchmt), 8, 539, 2007. 9. Kocatürk, T. et al. Post-LASIK epithelial dendritic defect associated with Alternaria. Cornea, 26, 1144, 2007. 10. Lo Cascio, G. et al. Utility of molecular identification in opportunistic mycotic infections: a case of cutaneous Alternaria infectoria infection in a cardiac transplant recipient. J. Clin. Microbiol., 42, 5334, 2004. 11. Romano, C. et al. Subcutaneous alternariosis. Mycoses, 48, 408, 2005. 12. Robertshaw, H. and Higgins, E. Cutaneous infection with Alternaria tenuissima in an immunocompromised patient. Br. J. Dermatol., 153, 1047, 2005. 13. Uenotsuchi, T. et al. Cutaneous alternariosis with chronic granulomatous disease. Eur. J. Dermatol., 15, 406, 2005. 14. Gallelli, B. et al. Skin infection due to Alternaria species in kidney allograft recipients: report of a new case and review of the literature. J. Nephrol., 19, 668, 2006. 15. Luque, P. et al. Treatment of cutaneous infection by Alternaria alternata with voriconazole in a liver transplant patient. Transplant. Proc., 38, 2514, 2006. 16. Vieira, R. et al. Cutaneous alternariosis in a liver transplant recipient. Rev. Iberoam. Micol., 23, 107, 2006. 17. Nudens, E. et al. Alternaria infectoria phaeohyphomycosis in a renal transplant patient. Med. Mycol., 44, 379, 2006.

Molecular Detection of Foodborne Pathogens 18. Ara, M. et al. Relapse of cutaneous Alternaria infectoria in a renal transplant recipient after 2 years. Acta Derm. Venereol., 86, 154, 2006. 19. Sood, N. et al. Subcutaneous phaeohyphomycosis caused by Alternaria alternata in an immunocompetent patient. Int. J. Dermatol., 46, 412, 2007. 20. Brasch, J., Busch, J.O. and de Hoog, G.S. Cutaneous phaeohyphomycosis caused by Alternaria infectoria. Acta Derm. Venereol., 88, 160, 2008. 21. Garau, J. et al. Alternaria osteomyelitis. Ann. Intern. Med., 86, 747, 1977. 22. Logrieco, A. et al. Epidemiology of toxigenic fungi and their associated mycotoxins for some Mediterranean crops. Eur. J. Plant Pathol., 109, 645, 2003. 23. Stinson, E.E. et al. Mycotoxin production by Alternaria species grown on apples, tomatoes, and blueberries. J. Agric. Food Chem., 28, 960, 1980. 24. Nishimura, S. and Kohmoto, K. Host-specific toxins and chemical structures from Alternaria species. Annu. Rev. Phytopathol., 21, 87, 1983. 25. Robiglio, A.L. and Lopez, S.E. Mycotoxin production by Alternaria alternata strains isolated from red delicious apples in Argentina. Int. J. Food Microbiol., 24, 413, 1995. 26. Li, F.G. and Yashizawa, T. Alternaria mycotoxins in weathered wheat from China. J. Agic. Food Chem., 48, 2920, 2000. 27. Nielsen, K.F. and Smedsgaard, J. Fungal metabolite screening: database of 474 mycotoxins and fungal metabolites for de-replication by standardised liquid chromatography-UV detection-mass spectrometry methodology. J. Chromatogr. A, 1002, 111, 2003. 28. Azcarate, M.P. et al. Alternaria toxins in wheat during the 2004 to 2005 Argentinean harvest. J. Food Prot., 71, 1262, 2008. 29. Schell, W.A. New aspects of emerging fungal pathogens. Clin. Lab. Med., 15, 365, 1995. 30. Laumaillé, C. et al. Cutaneous Alternaria infectoria infection after liver transplantation. Ann. Pathol., 18, 1998. 31. Mayser, P., Nilles, M. and de Hoog, G.S. Case report. Cutaneous phaeohyphomycosis due to Alternaria alternate. Mycoses, 45, 338, 2002. 32. Simmons, E.G. Alternaria themes and variations (236–243) Host-specific toxin producers. Mycotaxon, 70, 325, 1999. 33. Zur, G. et al. Development of a polymerase chain reactionbased assay for the detection of Alternaria fungal contamination in food products. J Food Prot., 62, 1191, 1999. 34. Asturias, J.A. A sensitive two-site enzyme-linked immunosorbent assay for measurement of the major Alternaria alternata allergen Alt a 1. Ann. Allergy Asthma Immunol., 90, 529, 2003. 35. Halaby, T. et al. Phaeohyphomycosis caused by Alternaria infectoria in a renal transplant recipient. J. Clin. Microbiol., 39, 1952, 2001. 36. Vartivarian, S.E., Anaissie, E.J. and Bodey, G.P. Emerging fungal pathogens in immunocompromised patients: classification, diagnosis, and management. Clin. Infect. Dis., 17, S487, 1993. 37. Andersen, B., Krøger, E.G. and Roberts, R. Chemical and morphological segregation of Alternaria arborescens, A. infectoria and A. tenuissima species-groups. Mycol. Res. 106, 170, 2002. 38. Guillemette, T., Iacomi-Vasilescu, B and Simoneau, P. Conventional and real time PCR-based assay for detecting pathogenic Alternaria brassicae in Cruciferous seed. Plant Dis., 88, 490, 2004.

Alternaria 39. White, T.J. et al. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: PCR Protocols: A Guide to Methods and Applications, pp. 315–322. Innis, N., Gelfand, J. and White, T. (eds.). Academic Press, Inc., New York, NY, 1990. 40. Haugland, R.A., Heckman, J.L. and Wymer, L.J. Evaluation of different methods for the extraction of DNA from fungal conidia by quantitative competitive PCR analysis. J. Microbiol. Methods, 37, 165, 1999. 41. Ferrer, C. et al. Rapid molecular diagnosis of posttraumatic keratitis and endophthalmitis caused by Alternaria infectoria. J. Clin. Microbiol., 41, 3358, 2003. 42. Ferrer, C. et al. Detection and identification of fungal pathogens by PCR and by ITS2 and 5.8S ribosomal DNA typing in ocular infections. J. Clin. Microbiol., 39, 2873, 2001. 43. Millar, B.C. et al. Isolation of Alternaria alternata from an emollient cream: implications for public health. Mycopa­ thologia, 156, 273, 2003. 44. Hong, S.G., Liu, D. and Pryor, B.M. Restriction mapping of the IGS region in Alternaria spp. reveals variable and conserved domains. Mycol. Res., 109, 87, 2005. 45. Dubois, D. et al. Cutaneous phaeohyphomycosis due to Alternaria infectoria. Mycopathologia, 160, 117, 2005. 46. Kusaba, M. and Tsuge, T. Phylogeny of Alternaria fungi known to produce host-specific toxins on the basis of variation in internal transcribed spacers of ribosomal DNA. Curr. Genet., 28, 491, 1995. 47. Laich, F.S. et al. Molecular characterization of Alternaria alternata causing ocular infection: detection of IGS-RFLP intraspecific polymorphism. Med. Mycol., 12, 1, 2008.

527 48. Roberts, R.G., Reymond, S.T. and Andersen, B. RAPD fingerprint analysis and morphological segregation of small-spored Alternaria species and species groups. Mycol. Res., 104, 151, 2000. 49. Iram, S. and Ahmad, I. Analysis of variation in Alternaria alternata by pathogenicity and RAPD study. Pol. J. Microbiol., 54, 13, 2005. 50. Johnson, R.D. et al. Cloning and characterization of a cyclic peptide synthetase gene from Alternaria alternata apple pathotype whose product is involved in AM-toxin synthesis and pathogenicity. Mol. Plant Microbe Interact., 13, 742, 2000. 51. Andersen, B. et al. Real-time PCR quantification of the AM-toxin gene and HPLC qualification of toxigenic metabolites from Alternaria species from apples. Int. J. Food Microbiol., 111, 105, 2006. 52. Johnson, R.D. et al. A polymerase chain reaction-based method to specifically detect Alternaria alternata apple pathotype (A. mali) the causal agent of Alternaria blotch of apple. Phytopathology, 90, 973, 2000. 53. Tournas, V.H., Kohn, J.S. and Katsoudas, E.J. The identification of antibacterial compounds for the development of enhanced media for the detection of foodborne fungi. Int. J. Food Microbiol., 118, 83, 2007. 54. Haugland, R.A., Brinkman, N. and Vesper, S.J. Evaluation of rapid DNA extraction methods for the quantitative detection of fungi using real-time PCR analysis. J. Microbiol. Methods, 50, 319, 2002. 55. Akamatsu, H. et al. Molecular karyotypes for Alternaria plant pathogens known to produce host specific toxins. Curr. Genet., 35, 647, 1999.

38 Aspergillus

Giancarlo Perrone, Antonia Gallo, and Antonia Susca Institute of Sciences of Food Production

Contents 38.1 Introduction.................................................................................................................................................................... 529 38.1.1 Classification and Taxonomy of Major Foodborne Aspergillus....................................................................... 530 38.1.1.1 Section Flavi.....................................................................................................................................531 38.1.1.2 Section Circumdati...........................................................................................................................531 38.1.1.3 Section Nigri.................................................................................................................................... 532 38.1.2 Biology, Ecology, and Toxicity of Major Foodborne Aspergillus..................................................................... 532 38.1.2.1 Aspergillus Ochratoxin A Producing Species................................................................................. 532 38.1.2.2 Aspergillus Aflatoxin Producing Species........................................................................................ 533 38.1.3 Current Laboratory Diagnostic Techniques for Aspergillus............................................................................. 534 38.1.3.1 Ochratoxin A Producing Aspergillus Species................................................................................. 535 38.1.3.2 Aflatoxin Producing Aspergillus Species........................................................................................ 537 38.2 Methods.......................................................................................................................................................................... 539 38.2.1 Molecular Detection of Ochratoxigenic Aspergillus Species in Food.............................................................. 539 38.2.1.1 Coffee............................................................................................................................................... 539 38.2.1.2 Grapes.............................................................................................................................................. 540 38.2.2 Molecular Detection of Aflatoxigenic Aspergillus Species in Food................................................................. 541 38.2.2.1 Figs................................................................................................................................................... 541 38.2.2.2 Corn................................................................................................................................................. 541 38.2.2.3 Peanut Kernel................................................................................................................................... 542 38.2.2.4 Wheat............................................................................................................................................... 542 38.3 Conclusions and Future Perspectives............................................................................................................................. 543 References.................................................................................................................................................................................. 545

38.1  Introduction The first description of Aspergillus dated from 1729, when P. A. Micheli named an organism with characteristic sporebearing structure resembling an aspergillum (a device used by the Catholic church to sprinkle holy water) as Aspergillus. The genus Aspergillus, a member of the phylum Ascomycota, is among the most abundant and widely distributed organism on earth; it includes over 250 known species and it is one of the most economically important of the fungal genera. Aspergillus spp. are widespread geographycally and can be either beneficial or harmful depending on the species and on the substrate being utilized. To adapt to the variety of niches they inhabit, they have evolved a myriad of metabolites. Some of these have been exploited by humankind.1,2 A number of Aspergillus-related patents have been issued for medical compounds, for example lovastatin, produced by A. terreus, was one of the first commercially successful cholesterol-lowering drugs.3 A number of antibiotic, antitumoral, and antifungal agents have been derived from Aspergillus metabolites. Strains of Aspergillus are diffusely used in industrial production like soy sauce (A. oryzae), several organic acids and enzymes (A. niger, A. aculeatus, A. carbonarius). Two of the

most important industrial products produced by Aspergilli are amylase and citric acid.4 They are common soil saprophytes which play an essential role in recycling carbon and nitrogen from organic debris.5 Unfortunately, the positive economic impact of the metabolites produced by this genus is more than balanced by negative aspects, in fact Aspergilli are one of the major causes of degradation of agricultural products. Although they are not considered to be major causes of plant disease, Aspergillus species are responsible for several disorders in various plants and plant-products; mainly A. niger, A. flavus, and A. fumigatus species also cause animal and human disease. Due to their importance in medicine, biotechnology, and foods, Aspergilli are in the spearhead studies concerning various aspects of fungi. Differentially from the common specialized plant pathogens like rust, powdery mildew and some Fusarium species, Aspergillus species are opportunistic pathogens without host specialization, and frequently isolated as food contaminants. Only a limited number of Aspergillus species are able to invade living plant tissues, while most of the species are found as storage mould on plant products. They can contaminate agricultural products at different stages including pre- and post-harvest, processing, and 529

530

Molecular Detection of Foodborne Pathogens

A

B

Figure 38.1  (A) Aspergillus flavus on nut. (B) Aspergillus carbonarius on grape berries.

handling (Figure 38.1). Changes due to spoilage by Aspergillus species can be of sensorial, nutritional and qualitative nature like pigmentation, discoloration, rotting, development of offodors, and off-flavors. Moreover, a number of pathogenic and saprophytic species produce toxigenic secondary metabolites (mycotoxins) on host tissue and plant products, so the most notable consequence of their presence is mycotoxin contamination of foods and feeds. The main mycotoxins produced by species belonging to Aspergillus genus are: aflatoxins (B1, B2, G1, G2); ochratoxin A; patulin; sterigmatocystin; cyclopiazonic acid; penicillic acid; citrinin; cytochasalin E; verruculogen and fumitremorgin A and B. Among these, the most important are the aflatoxins and ochratoxin A.6 Aflatoxins B1, B2, G1, G2 (AFs) are the most toxic and carcinogenic naturally occurring mycotoxins. Due to their extreme hepatocarcinogenicity, extensive research has been carried out on the natural occurrence, identification, characterization, biosynthesis, and genetic regulation of aflatoxins.7–9 Aflatoxins pose a risk to human health because of their extensive pre-harvest contamination of corn, cotton, soybean, peanuts, and tree nuts, and because residues from contaminated feed may appear in milk. Several aflatoxin outbreaks in humans after consumption of contaminated grains have been documented.10,11 The most important aflatoxin producing species belong mainly to Aspergillus Section (Sec.) Flavi, including A. flavus, A. parasiticus and several other species.7 Ochratoxin A (OTA) is a potent nephrotoxin which may contaminate various food and feed products (grains, legumes, coffee, dried fruits, beer, and wine, and meat). It also exhibits carcinogen, teratogen and immunotoxic properties in rats and possibly in humans.12 Studies on genotoxicity of OTA remain controversial.13 OTA is receiving increasing worldwide attention because of its wide distribution in food

and feed and of the risk for human health due to low levels of OTA contamination of a wide range of different foods.14 The economically most important OTA producers belong to Aspergillus sections Circumdati and Nigri.15,16 In this chapter we summarize the main aspect of the morphology, ecology, epidemiology, toxigenicity, with a particular emphasis on the diagnostic methods relevant to the most important sections of Aspergillus foodborne pathogen namely: Sec. Flavi, Circumdati, and Nigri occurring in several agricultural products and responsible of aflatoxin and ochratoxin A contamination of food and feed.

38.1.1 Classification and Taxonomy of Major Foodborne Aspergillus Aspergillus is a complex genus which comprises more than 250 recognized species covering 12 distinct teleomorph genera, all of which are members of family Trichocomaceae of the order Eurotiales. Various rearrangements of the species groups, sections and subgenera were made in Aspergillus over the course of last two decades due to the tremendous amount of genetic studies and sequences published; however little progress has been made regarding the understanding of the relationship among its subgeneric groups. Recently, based on phylogenetic analysis of sequence data using three different genes, the genus Aspergillus has been divided into eight subgenera which comprise 23 different Sections.17 These subgenera included five out of six identified by Gams et al.18 and three new subgenera, while the subgenus Clavati, previously described,18 was included in the Fumigati one. In the classification of the species and in the recognition of the genus, the main morphological feature is the “aspergillum,” consisting of a conidiophore stipe terminating in a vesicle, which in

Aspergillus

turns bears either a single palisade of cells (phialides, primary sterigmata) giving rise to conidia, or two palisades of cells, with those arising from the vesicle and one or two layers of synchronously formed specialized cells (phialides) and asexually formed spores called conidia. An Aspergillus with phialides borne directly on the vesicle is referred to as a “uniseriate” Aspergillus. If a second layer of specialized cells is present on the vesicle, the Aspergillus is referred to as “biseriate”. The second layer of cells, located between the phialides and vesicle, are called metulae. The Aspergillus stipe is often thick walled and has no or only a few thin septa. The basal part of the stipe is called “foot-cell,” which is, actually, not a separate cell, and it is often typical and useful for recognizing the conidiogenous structures of the genus. The morphological keys to common mycotoxigenic and foodborne Aspergillus species are based on macromorphology features like colors of mycelium and conidia, reverse color, colony diameter on different culture media and growth temperatures, presence of exudates, sclerotia and cleistothecia; and on micromorphology features like heads radiate or columnar, uniseriate or biseriate, size and shape of vesicles (spherical, spathulate, etc.), size and roughness of conidia, presence of Hülle cells and shape, presence of ascospores, ornamentation, color. Taxonomic and synoptic keys to Aspergillus species identification are available on various books and manuals.19–21 In foods, commonly encountered species are from the following subgenera and relevant sections: Subgenera: • • • • •

Aspergillus: Sec. Aspergillus, Restricti Fumigati: Sec. Fumigati, Clavati Nidulantes: Sec. Nidulantes, Ochraceorosei Circumdati: Sec. Flavi, Nigri, Circumdati, Candidi Terrei: Sec. Terrei

Among these, subgenus Circumdati is the most important source of food spoilage and mycotoxins accumulation in food, unless also the subgenus Aspergillus causes most spoilage problems in stored and processed foods. In fact, species economically important for agri-food productions belong mainly to the sections Circumdati, Flavi, and Nigri; with the exception of some minor important species belonging to the sections Nidulantes and Ochraceorosei. In particular, the most common species are A. niger and A. flavus, followed by A. parasiticus, A. ochraceus, A. carbonarius, A. nomius, A. alliaceus (Petromyces alliaceus), and recently also A. westerdijkiae, become important in coffee.6,16 Briefly herein are summarized the main morphological traits of the most important Aspergillus foodborne species. 38.1.1.1 Section Flavi It includes species with conidial heads in shades of yellowgreen to brown, and dark sclerotia, and comprises 23 species and subspecies according to the phylogenetic analysis of ITS and 28S rDNA sequences.22,23 A. flavus is the main species of the section due to its distribution and aflatoxigenicity. It is characterized by bright

531

yellow-green colonies (sometimes yellow), most heads have metulae and phialides, heads are radiate, conidia are smooth to finely roughened and of variable size (globose to ellipsoidal). It grows rapidly at 37°C. A. nomius has conidia similar to A. flavus, small and elongated (bullet-shaped) sclerotia, and may be distinguished by production of both B and G aflatoxins. The close rare species A. bombycis could be distinguished by its slow growth at 37°C and smooth stipe walls; also it may produce both aflatoxins. A. parasiticus colonies are dark green (never yellow), heads usually have only phialides (metulae occasionally), conidia are rough walled and usually more uniform in size than A. flavus. A parasiticus comprises a higher percentage of toxigenic isolates, producing both aflatoxins B and G. A. toxicarius has been synonymized to A. flavus24 and is closely related to A. parasiticus. It is still difficult to distinguish both species and its species definition is not completely resolved.25 A. tamarii and A. pseudotamarii, similar and closely related to A. flavus, also show rapid growth at 37°C, they may be distinguished by colonies more brown than A. flavus and by conidia very rough to tuberculate, much rougher than A. flavus or A. parasiticus. A. pseudotamarii differs from A. tamarii in production of aflatoxins (B type). The recently described aflatoxin producing species A. parvisclerotigenus and A. minisclerotigenes are difficult to differentiate morphologically from A. flavus, and A. ara­ chidicola from A. parasiticus. However, A. parvisclerotigenus and A. minisclerotigenes have both smaller sclerotia than A. flavus strains and produce both aflatoxins (B and G type), while they differ from each other only by molecular data and sclerotia extrolites production. A. arachidicola instead, is more similar to A. parasiticus and differs by a less dark green color of the culture, more biseriate conidiophores and the production of chrysogine.26 38.1.1.2 Section Circumdati It includes species with biseriate conidial heads in shades of yellow to ochre, important for the production of several mycotoxins among which ochratoxin A, named after the producer A. ochraceus; this section comprises 22 species of which seven have been recently identified.16,17 A. ochraceus was the main important species of this section relevant for the contamination of food by ochratoxin A, until recently two new closely related species, A. westerdijkiae and A. steynii, were characterized from strains previously identified as A. ochraceus and OTA producers.16 The main morphological features of these three important species are: colonies ochre or pale yellow; large radiate and biseriate heads; closely packed metulae and small phialides, smooth or finely roughed conidia. A. ochraceus has a variable growth at 37°C, and many isolates form pinkishbrown sclerotia; A. westerdijkiae differently is a strong producer of ochratoxin A (possibly the main source of this toxin from Sec. Circumdati), it doesn’t grow at 37°C, and the sclerotia are white/cream; A. steynii doesn’t grow at 37°C and has broadly ellipsoidal conidia “en masse” with pale

532

yellow color on MEA (Malt extract agar). The three species are very difficult to differentiate without the support of molecular and biochemical data. Many other species closely related to A. ochraceus have been recently described in Sec. Circumdati, but they are apparently rare and may not be important as potential mycotoxins contamination in foods and beverages.27 38.1.1.3 Section Nigri Named also “black aspergilli,” they have a significant impact on modern society as they cause food spoilage, and are used in the fermentation industry. This Section includes species with biseriate or uniseriate conidial heads in shades of brown-violet to black, and sclerotia with different color and size in 14 out of 19 species considered. The taxonomy of the Section is still not completely resolved, especially within the A. niger species aggregate (a group of morphologically indistinguishable species), often leading to misidentification of the species distribution in food. The recent revision of the taxonomy reveals that there are 19 taxa, with five of these producing ochratoxin A,28 however A. niger is the most frequently reported species in food together with A. carbonarius, A. japonicus and A. aculeatus. A. niger is characterized by its very dark brown to black colonies, conidial heads are radiate and biseriate with wide and spherical vesicles, conidia are globose irregularly roughened with ridges and bars. Optimal growth conditions are 35–37°C. However, A. niger group comprises an aggregate of at least four to seven species morphological similar or indistinguishable; of these the most frequent isolated are A. niger and A. tubingensis, while A. foetidus and A. brasiliensis are detected at lower frequencies, and the new described species A. piperis, A. lacticoffeatus, A. costaricaensis are rarely found in food commodities.15,29 A. carbonarius has optimal growth at 32–35°C and it can easily be distinguished from other biseriate species due to its big, spiny conidia; and the stipes up to several millimeters long. A high percentage of strains of this species (98–100%) have been shown to produce OTA. Other biseriate species similar to A. carbonarius, but difficult to distinguish, are A. ibericus, with light smaller conidia than A. carbonarius and no OTA production, and the producing species A. sclerotioniger with yellow mycelium, orange to brown sclerotia but smooth to verruculose conidia.15,30 Among the uniseriate species of the Section Nigri, A. japonicus and A. aculeatus are the most common isolated from food, but they are not distinguishable by morphology. They have uniseriate heads, conidia usually rough, varying from subglobose to ellipsoidal and echinulate with evenly spaced spines. A. aculeatus has larger conidial heads and conidia more ellipsoidal in shape. Recently a new closely related species isolated from grapes, A. uvarum, has been described. This is more similar to A. japonicus, it grows more slowly at 37°C than A. japonicus or A. aculeatus; these species are distinguished only by molecular and biochemical data.31

Molecular Detection of Foodborne Pathogens

38.1.2 Biology, Ecology, and Toxicity of Major Foodborne Aspergillus Aspergillus species have been described as ubiquitous, and this term could be applied both to substrate utilization as well as to geographic distribution; however in subtropical and tropical regions their occurrence is more common than the Penicillium species. As underlined previously the main species of Aspergillus occurring on food products belong to the subgenus Circumdati. Particular attention has been posed in the last decades to the toxigenic species due to the risks for humans and animals exposed by the consumption of food and vegetable products contaminated by Aspergillusmycotoxins (e.g., aflatoxins and ochratoxin A). In this respect this chapter has been divided in two sections relevant to the Aspergillus foodborne ochratoxin and aflatoxin producing species, respectively (Table 38.1). 38.1.2.1  Aspergillus Ochratoxin A Producing Species Ochratoxins are produced mostly by Penicillium species in colder temperate climates, and by a number of Aspergillus species in warmer and tropical parts of the world. Aspergillus isolates usually produce both ochratoxin A and B (dechlorinated analogue of OTA), while Penicillium produce only OTA. Despite that Aspergillus and Penicillium are the two ochratoxigenic fungal genera, the genus Aspergillus contains the greatest number of potential producers, which belong mainly to Sec. Circumdati, with some important species in Sect. Nigri and Flavi. In Aspergillus Sec. Circumdati,

Table 38.1 Ochratoxigenic and Aflatoxigenic Foodborne Aspergilli Aspergillus species producing ochratoxin A Sec. CIRCUMDATI A. cretensis A. flocculosus A. ochraceus A. pseudoelegans A. roseoglobulosus A. sclerotiorum A. steynii A. sulphureus A. westerdijkiae Neopetromyces muricatus

Sec. FLAVI A. albertensis A. alliaceus

Sec. NIGRI A. carbonarius A. lacticoffeatus A. niger A. sclerotioniger A. tubingensis

Aspergillus species producing aflatoxins (B and G type) Sec. FLAVI A. arachidicola (AFB and AFG) A. bombycis (AFB and AFG) A. flavus (AFB) A. minisclerotigenes (AFB and AFG) A. nomius (AFB and AFG) A. parasiticus (AFB and AFG) A. parvisclerotigenus (AFB and AFG) A. pseudotamarii (AFB)

Sec. OCHRACEOROSEI A. ochraceoroseus (AFB) A. rambelli (AFB) Sec. NIDULANTES Emericella astellata (AFB) Emericella venezuelense (AFB)

Aspergillus

A. cretensis, A. flocculosus, A. pseudoelegans, A. roseoglobulosus, A. westerdijkiae, A. sulphureus, and Neopetromyces muricatus produce consistently high amounts of OTA.16 Two other species, A. ochraceus and A. sclerotiorum produce variable amounts of OTA, but less consistently, while four further species produce OTA inconsistently and in trace amounts according to the literature: A. melleus, A. ostianus, A. petrakii, and A. persii. The most important species regarding potential OTA contamination of food products like coffee, rice, beverages, and other foodstuffs are A. ochraceus, A. westerdijkiae and A. steynii.27 In Sec. Nigri, A. carbonarius is the higher ochratoxigenic species responsible for OTA contamination of grapes and by-products,32,33 other less important producing species are A. niger and A. tubingensis,34,35 and the rarely occurring A. lacticoffeatus and A. sclerotioniger.15 Recently also some strains belonging to uniseriate species, A. aculeatus and A. japonicus were reported as OTA producers, but these data need to be confirmed by molecular identification of the species.32,36,37 In Aspergillus Sec. Flavi, A. albertensis and A. alliaceus (teleomorph Petromyces) were found to produce high amounts of ochratoxins, and are considered to be responsible for ochratoxin A contamination of figs.38 Several of these species are synonymous or poorly defined and, in this respect, few of these species are known to contaminate foods with ochratoxin A, which complicates analysis of ochratoxin contamination of food and crops. However, because few geographic areas have been studied, because many fungi are capable of producing ochratoxins, and because of taxonomic problems in Sec. Circumdati and Nigri, it is not always clear which species are responsible for ochratoxin contamination of crops and food commodities.38,39 Various ecophysiological studies have been made to identify conditions (incubation temperature, water activity, pH, different substrates) favoring growth, sporulation and toxin production by potential ochratoxigenic species. However, depending on the criteria used for species identification and laboratory tests used, some data results can be confusing or controversial. In general, A. ochraceus, A. westerdijkiae and A. steynii, previously identified mainly as A. ochraceus, are reported as saprophytic storage fungi, growing between 8 and 37°C, with the optimum at 24–31°C, and optimal 0.95–0.99 aw, while the lower limit for growth is 0.79 aw on media containing sugars and 0.81 aw on media based on NaCl. A. ochraceus species group grows slowly at pH 2.2 and well between pH 3 and 10. The optimum of OTA production by these species is very variable on the basis of the substrate (corn, rice, coffee, grapes, etc.) from 15 to 35°C and 0.90–0.99 aw, while the minimal aw for OTA production is 0.80–0.85 depending on the substrate; this is the most important CCP in the storage of food and feed.40 Regarding Sec. Nigri, the ecophysiology of this group of fungi has been recently investigated due to the risks that they pose in grape and by-product contamination; the ecological and toxicological differences between these species are summarized below. Grape-derived isolates belonging to the A. niger species aggregate, comprise four different species not distinguishable by morphological characteristics.

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The most frequent species isolated from grapes are A. niger and A. tubingensis with a low percentage of OTA producing strains (5–10%), while A. foetidus and A. brasiliensis are detected at lower frequencies and are no OTA producers.29 Optimal growth conditions are 35–37°C and 0.93–0.98 aw (min 6–8°C and max 47°C) for these isolates. This is one of the most common group of species in a wide range of fresh and dry fruits, cereals, etc., and its enzymes and organic acids are used in food processing as “GRAS” (generally regarded as safe). OTA production by A. niger species aggregate normally occurs at 20–25°C and 0.95–0.98 aw.41 A. carbonarius, the main species responsible for OTA accumulation in grapes, and recently found also as a contaminant of cocoa and coffee,42,43 has optimal growth conditions at temperatures between 25 and 35°C (min 10°C and max 42°C) and 0.95–0.98 aw. Optimal conditions for OTA production by A. carbonarius are at 20°C and 0.95–0.98 aw.44 No OTA production is detected at 0.90 aw, some reports observed higher OTA production at 25°C.45 Among the uniseriate group, A. aculeatus, A. japonicus and the new A. uvarum, isolated from grapes, coffee, and animal feeds, have been occasionally reported, but never confirmed, as OTA producers. They grow at similar temperature as A. carbonarius with optimal conditions at 32–35°C and 0.95–0.98 aw. The ability to grow at reduced aw is better and faster than A. carbonarius, but lower in respect to A. niger aggregate. Regarding the Sec. Flavi species OTA producing A. alliaceus and A. albertensis are considered widely distributed, but not common and never identified as contributors of OTA contamination of vegetable products or food; only A. alliaceus has been rarely isolated in figs and tree nuts in California.38 About their ecophysiology no data are available, except information about a higher toxigenicity in respect to A. ochraceus and A. melleus species, which are the dominant species in fig orchards. As a consequence of this wide distribution of OTA producing Aspergillus species in three Sections, it is unclear in some cases which fungi are responsible for OTA contamination of food and the list of foods which can be contaminated has continuely expanded.46 In fact, OTA is found in cereals and cereal products, beans, groundnuts, oilseed, spices, dried fruits, coffee, beer, wine, and grape juice from many countries.46 In general, studies have revealed a tremendous intraspecific variation in OTA production due to a combination of phenotypic plasticity and genetic variation. The fungi responsible for food contamination vary from crop to crop and from place to place, therefore, correct identification of the potential ochratoxigenic species in food is important. 38.1.2.2  Aspergillus Aflatoxin Producing Species The diversity of ecological niches occupied by members of Aspergillus Sec. Flavi and the ability of some species to produce aflatoxin make this group of fungi one of the most studied to date. Species in this Section occur in nature as saprophytes in the soil and on decaying plant material or as parasites on plants, insects, and animals. Aspergillus Sec. Flavi

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species in general appear to be most abundant in subtropical and warm temperate regions, particularly in agricultural and desert soils, and decrease in density and species diversity with increasing latitude. The main important species are the so-called domesticated species, such as A. oryzae, A. sojae and A. tamarii used in various food fermentation processes, and the main aflatoxin producing species A. flavus and A. parasiticus. In fact, several species of this section are able to produce aflatoxins, but AFs in crop and food are mainly produced by A. flavus and A. parasiticus which coexist with and grow on almost any crop or food. In nature, A. flavus is one of the most abundant and widely distributed soil-borne molds, its optimal growth is at 28–37°C and 0.90–0.99 aw, but it can grows also at temperature from 12 to 48°C and aw 0.77.47 Optimum for production of AFs is 28–30°C and 0.99 aw, limit conditions are 15°C and 0.83 aw.48 A. flavus is a saprophytic fungus capable of surviving on many organic nutrient sources, however it is also a weak opportunistic pathogen of many agricultural crops such as corn, cotton, peanuts, tree nuts etc. Its significance as a plant pathogen is due to aflatoxin production in the seeds of several crops both before and after harvest, causing a health hazard for animals and humans. It is also considered the second leading cause of invasive and noninvasive aspergillosis in immunosuppressed patients after A. fumigatus. In fact, it can cause a broad spectrum of diseases in humans such us pulmonary aspergillosis, allergic bronchopulmonary aspergillosis, keratitis, pericarditis, etc.49 Not all A. flavus isolates produce AFs and those that do usually produce only B aflatoxins. The percentage of toxigenic A. flavus varies with strain, substrate, and geographic origin. Whereas A. flavus is broadly spread on soil and various crops, A. parasiticus is generally less abundant than A. flavus, and it infects primarily peanuts, while is uncommon in aerial crops. Almost all A. parasiticus isolates produce aflatoxins B and G. A. parasiticus needs a lower temperature for seed invasion than A. flavus and seems more adapted to soil survival, explaining its preference for peanuts with respect to A. flavus. A. parasiticus has similar temperature and aw limit as A. flavus for both fungal growth and AFs production, with aflatoxin production optimum between 24 and 30°C and high water activities (0.95–0.96).50 In general, ecology and epidemiology have been widely investigated for A. flavus and A. parasiticus with respect to the other Sec. Flavi species, which are mostly of minor importance for the agri-food industry. Both A. flavus and A. parasiticus populations overwinter in the soil as primary inoculum; drought stress and elevated temperatures promote infection in crops with consequent dispersal of aerial inoculum and successive accumulation of AFs in plant products.51 Population analyses in Sec. Flavi have evidenced great variability in morphological characters (phenotype) which renders the identification and discrimination of the species difficult, thus the presence of other aflatoxigenic species could be underestimated especially regarding the closely related species A. nomius and A. bombycis; and the newly described species A. minisclerotigenes, A. arachidicola, and A. parvisclerotigenus morphologically indistinguishable from A. flavus and A. parasiticus.

Molecular Detection of Foodborne Pathogens

In this respect the molecular and phylogenetic analysis of Sec. Flavi have evidenced three main clades: “A. flavus,” “A. tamarii” and “A. alliaceus,” and minor clades in which are scattered other aflatoxin-producing species such as A. nomius, A. bombycis, A. pseudotamarii. This probably indicates that the aflatoxin-producing ability was lost (or gained) several times during evolution. Another aflatoxin producing species, A. ochraceoroseus, was found to be unrelated to any of the species belonging to Sec. Flavi.52 This species, together with the recently described species, A. rambelli,25 are the only known to accumulate aflatoxin B1 and sterigmatocystin simultaneously and belong to Sec. Ochraceorosei, a sister group of Sec. Flavi. Additionally, AFs production has also been recently observed in two species outside of Sec. Flavi, Emericella venezuelense and E. astellata belonging to Aspergillus Sec. Nidulantes.25 Recent studies of genetic variability of A. flavus indicate that the name is currently used for a paraphyletic group of isolates that may produce aflatoxins B or G, and have large or small sclerotia. In particular, isolates with small sclerotia (group II) produce both aflatoxins B and G like A. parasiticus but they are clearly separated from it by molecular analysis, and deserve recognition as a new species,53 recently described as A. parvisclerotigenus.25 The other group (group I) includes isolates producing only aflatoxin B, and having large or small sclerotia, A. oryzae belongs to this group. It is interesting to note that although there are a lot of supporting data to classify A. oryzae and A. sojae as A. flavus and A. parasiticus respectively, as morphological variants, these taxa should be retained as separate species because of the regulatory confusion that conspecificity might generate in the food industry.54 Recently, two novel aflatoxin ­producing Aspergillus species, namely A. minisclerotigenes and A. arachidicola, were described from peanuts in Argentina. They are closely related to A. parvisclerotigenus and A. parasiticus, respectively.26 Nevertheless, the main cause of crop and food contamination by aflatoxins is due to A. flavus and secondary to A. parasiticus; the distribution, ecology and toxigenity of these newly emerging species should be taken into account by mycologists during surveys.

38.1.3 Current Laboratory Diagnostic Techniques for Aspergillus Fungi play an important role as spoilage or toxigenic microorganisms mainly of plant-derived foods. Besides their spoiling activity, many of the food-relevant fungi are able to produce mycotoxins. For most of these toxigenic fungi, molecular detection systems have been developed, while only a few methods have been developed for spoilage fungi. The fungal contamination of a food sample is usually assessed by classical mycological means, e.g., the plate count technique. It has the great advantage to detect a broad spectrum of fungal species and the disadvantage of time consumption, need for mycological expertise, and above all the possibility of misclassification, since morphological characters could be highly variable depending on the media and culture conditions. In this respect, control of the unwanted contaminations during

Aspergillus

large scaled bio-industrial fermentations necessitates fast and exact identification of the Aspergillus species. Therefore, considerable attempts have been carried out to develop fast and precise molecular tools, instead of laborious and time-consuming techniques. In particular, identification of Aspergillus species based only on morphological characteristics might be difficult; it usually requires more than one media and the implementation of selective media due to different competition abilities of the different species. In general, the direct plating technique or the dilution plating technique are applied as traditional methods for Aspergillus enumeration from food samples using Dichloran Rose Bengal Chloramphenicol agar (DRBC), preferably for fruits, vegetables, herbs, and fresh cereals and Dichloran 18% glycerol agar (DG18) for stored cereals, spices, and nuts.15 Selective media for identification of potential aflatoxigenic and ochratoxigenic strains have been also developed such as AFPA (A. flarus parasiticus agar) medium for detection of A. flavus and A. parasiticus by development of orange reverse pigment55 and CCA (coconut cream agar) for detection of aflatoxin and ochratoxin producing strains by UV fluorescence.56 Recently room temperature phosphorescence (RTP) was observed for aflatoxigenic strains grown on media commonly used in food mycology, to which methyl-βcyclodextrin plus bile salts (0.6% sodium deoxycholate) were added.57 However, these methods have a low degree of sensitivity, are difficult to standardize5,58 and because some fungi may be poorly characterized or because considerable expertise is required, misidentification can occur. In this respect, the use of DNA markers and PCR-based methods may provide useful tools to replace conventional methods to analyze either the taxonomic classification or even the detection of a mycotoxigenic potential isolate. There is no method (morphological, physiological or molecular) that works flawlessly in recognizing species. Traditional Aspergillus taxonomy, based on phenotypical characters only, gives mostly a satisfactorily delimitation of the taxa. In fact, in several sections of the genus, many morphological variations occur, resulting in debatable taxonomy schemes. Therefore, this is why Aspergillus taxonomists have recently embraced the polyphasic approach in order to examine the variability within the species.27 Most Aspergillus species have been defined on the basis of morphology, with the additional consideration of molecular and extrolite data used in recent years.16,31,59 Molecular phylogenetics has uncovered cryptic speciation in a number of taxa,54,60,61 suggesting that morphological characters provide a very broad species concept that does not reflect the true extent of evolutionary divergence and reproductive isolation, as appears to be the case in fungi.62 However, molecular polymorphisms provide the greatest number of variable characters for fungal taxonomy. They can be generated using a widely available technology, sustained by extremely well-developed bioinformatic infrastructure that allows worldwide communication and comparison of results. This utility ensures that molecular characters have a primary role in recognising fungal taxa.63 In this regard, a number of molecular techniques are being utilized in order to type and to understand the molecular relationship among

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Aspergillus strains, contributing with conventional taxonomy, to the definition of species in the Aspergillus genus and detecting individual species from crops, food, and feed not showing disease symptoms. Finally, the availability of protocols to isolate DNA from food samples is important in a PCR-required purity, as it is known that food components can interfere with the PCR reaction. This task becomes fundamental when applied to real-time PCR for the quantification of the inoculum (fungal biomass). If a diagnostic PCR is used to assess the safety of a food sample, false-negative reactions resulting from PCR inhibition must be recognizable, hence the importance of an internal standard.64 During the last decade, molecular analytical methods have been widely studied and developed for the identification and detection of foodborne toxigenic Aspergillus species. They are fixed on anonymous markers or conserved genes for ochratoxigenic species of which no specific target sequences of the OTA biosynthetic pathway are known. They are directed differently on AFs biosynthetic genes for the aflatoxigenic species, the aflatoxin biosynthetic gene pathway being widely elucidated.64 In this case, it has been possible to develop detection procedures able to differentiate between mycotoxin and non-mycotoxin producing strains by monitoring the mRNA expression levels of biosynthetic genes. These two aspects are summarized and discussed in the following sections. 38.1.3.1 Ochratoxin A Producing Aspergillus Species In general ochratoxigenic fungi responsible for food contamination vary from crop to crop and from place to place, and often are morphological and phenotypical indistinguishable from other Aspergillus species. For these reasons, in recent years a large amount of work has been done in developing molecular tools and strategies for the correct identification of the potential ochratoxigenic species in food. In this regard, the most promising techniques are PCR based such as random amplified polymorphic DNA (RAPD), microsatellite length polymorphisms (MLP), short tandem repeats (STR) and amplified fragment length polymorphism (AFLP), sequencing of rRNA genes and protein-coding gene, as well as methods based on noncoding repetitive sequences in combination with restriction fragment length polymorphisms (RFLP). Use of these and other methods has been reviewed by Varga.65 Three recent additions to this list are multilocus sequence typing (MLST),66 coding tandem repeats and retrotransposon insertion-site context (RISC) typing.67 The specificity of the PCR-based methods depends on the target gene and its taxonomic relations between species. If primers for diagnostic PCR are based on well analyzed DNA regions, which can differentiate mycotoxin producers and nonproducers, it might be possible to develop a PCR system which can predict mycotoxin producing capability of a given strain. Today, little information is known about the biosynthetic pathway in Aspergillus OTA producers.68,69 Therefore today the best way to assess Aspergillus OTA toxicological risk is correctly determining fungal species occurring on crops because each species is associated to a specific capability to produce and accumulate mycotoxins.

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The exact identification is very important to avoid overestimation of contamination and toxicological risk.34,35 Therefore, commonly species-specific primers have been designed in different studies using rRNA genes, calmodulin (cmdA) gene, or anonymous genomic markers from AFLP, cDNA AFLP and RAPD fingerprinting analyses. Recently, various species and gene specific PCR primers have been developed for detection and identification of OTA producers in conventional and real-time PCR applications.70 Schmidt et al.71,72 used AFLP to detect specific markers for A. ochraceus and A. carbonarius. The authors compared strains of both species isolated from coffee related sources and ­demonstrated that AFLP fingerprints distinguished well at the species level both taxa from the closely related species; primers designed from the marker sequenced enabled detection of both species. Fungaro et al.73 developed primers with high specificity to A. carbonarius, using a fragment obtained by RAPD analysis which differentiates A. carbonarius from the other species. Various PCR assays were developed by using nucleotide sequences of well-defined fungal genes, the first PCR was set up for the detection of A. niger using sequences from the 18S rRNA gene, but the resulting primers were genus specific for Aspergillus.74 Specific variation within the ITS-regions of the rRNA genes was used by Sugita et al.,75 as a source for primer design to develop species-specific assays for the detection of A. fumigatus, A. flavus, and A. niger, the most frequently occurring fungi in human pulmonary infections. Based on the alignment of a partial sequence in representative strains of A. carbonarius, A. japonicus, and A. aculeatus, Perrone et al.76 identified three variable regions in the calmodulin gene sequences, exploited to design specific primers and to set up a PCR assay for identification and detection of A. carbonarius and A. japonicus/A. aculeatus strains isolated from grapes in Europe. Recently, with the aim to develop molecular tools for discrimination of species within Sec. Nigri, González-Salgado et al.77 designed PCR assays based on ITS region sequence to detect and identify A. niger, A. japonicus, A. heteromorphus, and A. ellipticus. However, the assay could not distinguish strains of A. niger from A. tubingensis. Anyway, more recently Susca et al.78 set up an assay for the accurate identification of A. niger and A. tubingensis in pure cultures, based on differences found in the calmodulin (cmdA) gene, usable also for multiplex PCR assays. Similarly, Patino et al.79 developed two ITS-based assays specific for A. carbonarius and A. ochraceus. Although this strategy promises a high degree of specificity for the primers selected, it has the disadvantage of producing amplicons of equal length in both PCR assays, which renders them unsuitable for multiplex assay. Moreover the identification requires a further step, a digestion of the resulting PCR product with the restriction endonuclease RsaI, able to cut only A. niger fragment. As was discussed in the previous paragraphs, the recently reviewed taxonomy of the Aspergillus Sec. Circumdati evidenced the major involvement of A. westerdijkiae as cause of OTA contamination of food instead of A. ochraceus. In this respect, Morello et al.80 exploring the nucleotide sequence variability in a region of the β-tubulin

Molecular Detection of Foodborne Pathogens

gene, designed a specific assay for detecting and quantifying A. westerdijkiae in Brazilian coffee beans. In fact, they compared several isolates from Brazilian coffee bean samples, previously identified as A. ochraceus, and most of them (84%) were re-identified as A. westerdijkiae. Recently, Susca et al.81 developed a PCR-single-stranded conformational polymorphism (SSCP) screening method based on sequence variations contained in about 180 bp region of calmodulin gene. Using this approach, 11 species including A. brasiliensis, A. niger, A. tubingensis, A. foetidus, A. aculeatus, A. uvarum, A. japonicus, A. ellipticus, A. heteromorphus, A. carbonarius, and A. ibericus could be distinguished based on their different PCR-SSCP profiles. A low-complexity oligonucleotide microarray (OLISA) has also been developed based on oligonucleotide probes obtained from sequences of the calmodulin gene for the detection of black aspergilli (A. carbonarius, A. ibericus, and A. aculeatus/A. japonicus) from grapes.82 Recent advances in DNA-based techniques such as real-time PCR (rt-PCR) are providing new tools for fungal detection and simultaneous quantification. Like any other fungal quantification technique, rt-PCR has also some drawbacks due to presence of fungi in vegetable samples in different forms i.e., spores, young mycelial fragments with many nuclei or old mycelial fragments with only a few nuclei. Nevertheless, it allows the specific detection of fungi in a mixed population which is by far the main advantage of PCR. In addition, post-PCR manipulations (such as gel electrophoresis) are not required, thus providing detection protocols that are highly sensitive, reliable, and suitable for high throughput analysis.83 Regarding A. carbonarius, various rt-PCR assays have been developed using a TaqMan or SYBR-Green I approach targeting conserved regions of the A. carbonarius calmodulin gene,84 the acyltransferase domain85 or the β-ketosynthase (KS)86 of a polyketide synthase (PKS) gene. For DNA purification from grapes, the first two groups of authors use a combination of grinder homogenization step, a plant DNA extraction procedure and a column purification step, whereas last group combines pulsifier equipment for homogenization and a commercial kit. All of them obtained results in the same range of detection limit, although the TaqMan rt-PCR system developed by Mulè et al.84 and Selma et al.86 has a detection limit about 1 log order lower (more sensitive) than the detection limit reported for the SYBR-Green I system used by Atoui et al.85 Both methods provide a good tool for early detection of this OTA-producing fungus in raw cultures such as grapes or coffee thereby preventing OTA from entering the food chain. In the case of grapes, the quantification of A. carbonarius at the early stage of maturation, could allow the growers to take necessary measures in order to reduce the final OTA contamination level in the field. A rt-PCR assay for quantitative estimation of A. ochraceus DNA is also available based on A. ochraceus specific primer pair, designed on anonymous DNA fragment.71,87 The possible use of genes involved in OTA production for PCR detection systems has been investigated, but it did not give reliable results.88 To date, although still incomplete, the

Aspergillus

most profound knowledge about genetics of the OTA biosynthetic pathway is available for P. nordicum. Geisen et al.89 detected and characterized the ­otapksPN gene, a polyketide synthase gene from OTA producing Penicillium spp. None of the sequences for OTA biosynthetic pathway genes identified in Penicillium nordicum90 has homologies with genes found in OTA producing Aspergillus spp. In particular, no homologies were found between sequences of the otapksPN gene and a polyketide synthase demonstrated to be potentially involved in OTA biosynthesis in A. ochraceus.68 Atoui et al.91 characterized the KS domain sequences of five different pks genes in A. carbonarius. However, none proved to be conclusively connected to the biosynthesis of toxin. Dao et al.92 sequenced the ketosynthase domain of a polyketide synthase gene in A. ochraceus, and designed two PCR assays, one detecting specifically A. ochraceus and the other identifying in a “group specific manner” fungi potentially able to produce OTA and the mycotoxin citrinin. The spectrum of species detected by the latter assay comprised A. carbonarius, A. melleus, A. ochraceus, P. citrinum, and P. verrucosum, but also Monascus ruber as a typical producer of citrinin in food and feed, among other Monascus spp. Detecting producers of both toxins may be an advantage because they can be found to be cooccurring in contaminated cereals.93 Finally, systems for molecular detection and identification of most of the fungal Aspergillus species known to produce OTA have been set up in the last few years. However, several new species have also been described, so future development must focus on PCR-based diagnostics for these new species and to improve their detection in food and commodities, because most of the detection procedure have been only tested on pure culture. 38.1.3.2 Aflatoxin Producing Aspergillus Species Due to their deleterious effects on human and animal health as well as their impact on international trade of agricultural commodities, the aflatoxin producing fungi have been the target of the first PCR-based diagnostic methods. The complete understanding of the biosynthetic pathway for aflatoxin production, achieved over 30 years of investigations, has led to the development of PCR assays aimed to detect the presence or the expression of the genes involved in the biosynthesis. About 25 genes and ORFs have been cloned and characterized which constitute the aflatoxin gene cluster within a 75-kb DNA region.8,94,95 In this chapter the new naming scheme, proposed by Yu et al.9 to promote the use of a uniform and conventional identification system, is adopted. It is based on the three-letter code “afl” followed by a capital letter to describe each individual gene confirmed to be or potentially involved in aflatoxin biosynthesis; when the genes are mentioned for the first time, the old names are given in brackets. In 1996, Geisen96 developed a multiplex PCR using primers for the three structural genes aflD (nor-1), aflM (ver-1) and aflP (omt-A). The multiplex PCR method, in which

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specificity is increased because a number of genes can be detected in one reaction, allowed to observe the triplet banding pattern for isolates of A. flavus, A. parasiticus and A. versicolor, this latter species producing only sterigmatocystin, a toxic intermediate of the aflatoxin biosynthesis. The system appeared to distinguish between aflatoxigenic Aspergillus species and other nonaflatoxigenic food related species, while it was less specific in differentiating between nonaflatoxin producing and aflatoxin producing strains of A. flavus. The pattern of negative strains of A. flavus varied and one of them showed all the three amplicons. A. sojae and A. oryzae, both of which are essentially identical with A. flavus and A. parasiticus, but typically do not produce aflatoxins, appeared to have sequences homologues to aflM and aflP while resulted lacking the aflD gene because of a sequence variation or a genic deletion. Applying a PCR diagnostic method targeting the same three genes, the expected amplicons were obtained from DNA isolated from infected figs, with uninfected figs ­giving no signals.97 The assay was also used for the detection of A. flavus in contaminated wheat.98 Among the genes involved in aflatoxin biosynthesis, there are two regulatory genes which control the level of structural genes expression: aflR and aflS. The protein AFLR, coded by the gene aflR, is supposed to bind the promoter region of most of the structural pathway genes activating gene expression; in addition, the aflR gene has an autoregulation function.99,100 The role of aflS is less characterized than that of aflR, but it seems to enhance transcription of the genes in some way.101 Shapira et al.102 proposed a diagnostic method to be applied in contaminated corn using the regulatory gene aflR (apa-2 in the paper) in addition to the aflM and aflP genes. Specific PCR products were obtained only with DNA from A. parasiticus with all three primer pairs, with the aflR primers failing to give a positive signal in A. flavus, likely for the subtle sequence differences of aflR gene between the two species. The aflR gene amplified from some nonaflatoxigenic strains of A. sojae presents differences that produce a nonfunctional protein and, therefore, they cannot produce aflatoxins. In addition, some strains of A. oryzae, even not being toxigenic, present an aflR gene, quite similar to the A. flavus aflR gene, which do not produce detectable transcript.103 It was postulated that when A. flavus strains have the A. oryzaetype aflR, they are not able to produce aflatoxins.104 Moreover some studies have identified a duplicated aflR gene in A. parasiticus, even though this second gene does not completely replace the first aflR gene.105 A multiplex PCR procedure using set of primers for four genes (aflR, aflD, aflM, and aflP) was developed by Criseo et al.106 The aflatoxigenic strains of A. flavus gave a quadruplet pattern indicating the presence of all the genes tested. Nonaflatoxigenic strains showed varying results, among which a complete pattern of four bands, which did not permit the discrimination with the aflatoxigenic strains. The same four genes were used in a multiplex PCR for the analyses of peanut kernels artificially contaminated with A. parasiticus and A. niger by Chen et al.107 The target DNA fragments were

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detected in the kernels infected with A. parasiticus and none was detected in the uninoculated or in the kernels infected with A. niger. Yang et al.108 applied successfully in vitro a multiplex PCR for the screening of potential aflatoxin-producing molds in Korean fermented food and grains, using as target the three aflatoxin biosynthetic genes aflI (avfA), aflP and aflM. A common problem encountered in PCR analysis is the interference caused by many product components such as polysaccharides, polyphenols, and other secondary compounds which may inhibit the activity of the polymerase, reducing sensitivity. This pitfall could be overcome by using adapted PCR or DNA preparation protocols. Manonmani et al.109 described an approach in which a nested PCR following a first PCR reaction, with very diluted DNA, didn’t show any negative or false-negative results due to the presence of food components. The PCR method, which had high sensitivity and specificity in detecting aflatoxigenic Aspergilli in selected food commodities, was based on the amplification of aflR sequence. Recently diagnostic methods based on the detection of messenger RNA (mRNA) transcripts for genes involved in mycotoxin production have being developed. They would give a clearer indication of mycotoxin production given that if these biosynthetic genes are being transcribed then the mycotoxin is being produced. However it should be considered that only some genes can be regarded as key genes, whose activation is directly coupled to the aflatoxin biosynthesis. The RT (reverse transcription)-PCR approach is useful to monitor the expression of specific genes under different physiological conditions allowing the study of the complex regulation of aflatoxin biosynthesis and its relationship to fungal development. Mayer et al.110 described the application of a real-time RT-PCR system for a quantitative monitoring of the expression of the aflD gene and its correlation to aflatoxin production by A. flavus in wheat samples. From the results, aflD mRNA could be detected about 48 h before the first produced aflatoxin B1 could be determined. The method allowed to determine quantitatively and very rapidly the infection in a food matrix by an aflatoxigenic Aspergillus and to monitor the process of aflatoxin B1 biosynthesis, resulting in a valuable tool for hazard analysis critical control point (HACCP) purposes. A real-time PCR system targeting the aflD gene to quantify the contamination of an aflatoxigenic A. flavus strain in plant-type foods like maize, pepper, and paprika had been applied by the same authors.111 A good correlation between the quantification of aflD and the cfu of the fungus was observed; however the aflD copy numbers were always higher. The analysis of a single gene expression may not be completely informative since the different genes of the biosynthesis pathway are subjected to complex regulation and interaction mechanisms, depending on development and physiological factors which have yet to be fully understood. More suitable are diagnostic RT-PCR systems based on the concurrent expression of more than one key gene involved in the biosynthetic pathway of aflatoxin, as a marker of the toxin production potential. Sweeney et al.112 employed an RT-PCR

Molecular Detection of Foodborne Pathogens

based detection system to monitor aflatoxin production and expression of two aflatoxin genes in A. parasiticus. Two primer pairs were designed to amplify the coding regions of the structural gene aflQ (ord1) and of the positive transcriptional regulator aflR. The technique allowed the correlation between the production of aflatoxin and the detection of the two specific gene transcription when A. parasiticus was cultured under permissive conditions. Significantly no aflQ or aflR transcripts were observed correlating with lack of aflatoxin production under nonpermissive conditions. However the functionality of the system was demonstrated in media but not in food matrix. In their work, Scherm et al.113 tested sets of primers matching the conserved regions of nine characterized structural genes (aflD, aflG, aflH, aflI, aflK, aflM, aflO, aflP, and aflQ) and the regulatory genes aflR and aflS to be used in RT-PCR. Three genes, namely aflD, aflO (omtB) and aflP, were identified which gave the best performance in the analysis. Their expression was always correlated to the aflatoxin production in inducing and noninducing media measured by fluorescence HPLC. The presence of the transcripts of these three genes allowed to distinguish between aflatoxin-producing and nonproducing strains of A. flavus and A. parasiticus. Detection of transcripts of other tested genes was not always correlated to the toxicity of the strains, in particular with the primers designed in this study aflQ and aflR did not result as useful targets, this differs from previous reports. The simultaneous detection of several specific mRNAs in a single RNA sample is possible with the combination of multiplex PCR and RT-PCR approaches, which other than being relatively simple and rapid, may reduce false-negative results. An example of the application of a multiplex RT-PCR method for the screening and discrimination of aflatoxin producing and nonproducing strains of A. flavus was reported by Degola et al.114 The procedure employed the two regulatory genes aflR and aflS and the structural genes aflD, aflO, and aflQ of the aflatoxin cluster of A. flavus and was applied to strains isolated from corn grains. While all the expected amplicons were detected in the genomic amplification of the aflatoxin producer strains, one or more of them, except the aflQ amplicon, were missing in most of the nonproducing strains, indicating a possible genic mutation. The multiplex RT-PCR analyses revealed a good correlation between gene expression of the five aflatoxin genes and aflatoxin production, except for one strain that apparently transcribed all the relevant genes but did not produce aflatoxin in the medium. From the analysis, the structural genes were expressed following the transcription of the regulatory genes aflR and aflS. The aflatoxin nonproducer isolates examined in this study failed to give a positive transcriptional signal for any of the genes. The PCR diagnostic methods illustrated here and based on biosynthetic genes were mainly developed for A. flavus and A. parasiticus, which are the major species of concern for production of aflatoxin in agricultural commodities and share a 90–99% identity in aflatoxin clustered genes.9 A PCR-based detection assay has not yet been described for

539

Aspergillus

the rarely encountered aflatoxin producers A. ­bombycis, A. ochraceoroseus and A. pseudotamari. A specific PCR detection of A. nomius using sequences from the ITS region of the rRNA gene was described by Haugland and Vesper115 together with a fluorescently labeled probe to be applied in a TaqMan quantitative real-time PCR assay.

38.2  Methods The availability of protocols to isolate fungal DNA from a food sample is an important aspect, especially for PCR applications. A low rate of DNA isolation and the presence of inhibitor compounds of DNA polymerase can interfere with results and can lead to down estimation or ­false-negative of fungal contamination. In particular, quantification of the fungal biomass is not an easy task. If one target gene is present in the genome, the number of copies reflects the genome number, which is proportional to the fungal biomass or cell number. In the case of filamentous fungi, the biomass consists of different cell types including vegetative spores, ascospores, and hypha. Spores can be uninucleate or multinucleate depending on the fungal species, whereas hyphal cells are multinucleate. PCR can detect dead cells, therefore copy numbers determined by real-time PCR are higher than the numbers of viable cells determined by the plate count ­method.111 In the following sections, some molecular detection methods for ochratoxigenic and aflatoxigenic Aspergillus foodborne fungi developed and tested directly on raw food matrices are described taking into account their specificity, robustness and reproducibility.

38.2.1 Molecular Detection of Ochratoxigenic Aspergillus Species in Food 38.2.1.1  Coffee Sample preparation. (1) Immerse the contaminated coffee beans in a 0.4% hypochlorite solution for 2 min.

Species A. carbonarius73

A. ochraceus71

A.niger117

Primers (5′–3′) OPX7F809: AGG CTA ATG TTGA TAA CGG ATG AT OPX7R809: GCT GTC AGT ATT GGA CCT TAG AG OCA-V: ATA CCA CCG GGT CTA ATG CA OCA-R: TGC CGA CAG ACC GAG TGG ATT OPX7F372: CAG TCG TCC AGT ACC CTA AC OPX7R372: GAG CGA GGC TGA TCT AAG TG

(2) Then rinse them with sterilized distilled water for 1 min. Repeat this washing procedure three times. (3) To extract the DNA, use a CTAB-based protocol.116 First, crudely grind one coffee bean using a hammer and then macerate to a fine powder with the addition of liquid nitrogen. Add to the resulting powder, 830 μl of extraction buffer (4% CTAB, 1.4 M NaCl, 20 mM EDTA, 100 mM Tris-HCl, 1% PVP) and 0.4% of β-mercaptoethanol. (4) Incubate the mixture at 65°C for 20 min, cool and keep at room temperature for 10 min, then add 600 μl of chloroform: isoamyl alcohol (24:1, v/v). (5) Centrifuge the resulting emulsion at 14,000×g for 7 min. Transfer the upper aqueous phase to a clean tube and repeat the deproteinization procedure as described above. (6) Add to the aqueous phase, 1 volume of isopropanol, mix and incubate at −20°C for 2 h. (7) Centrifuge the material for 5 min at 14000×g and rinse the pellet twice with 70% ethanol and leave to dry at room temperature. (8) Resuspend the pellet in 50 μl of TE buffer (10 mM Tris-HCl, 1 mM EDTA). (9) The total DNA obtained from one coffee bean was serial diluted 101, 102, and 103 fold and used as a template DNA for the multiplex PCR reaction Detection procedures. (i) Qualitative PCR for A. carbonarius, A. ochraceus, A. niger:117 (1) Prepare PCR mixture (25 μl) containing 3.0 μl DNA of each template, 0.5 μl MgCl2 (50 mM), 2.5 μl Taq DNA polymerase buffer 10×, 1.0 μl each primer (10 µM), 10.9 μl H2O, 0.1 μl Taq polymerase (5 U/μl) and 2.0 μl dNTP (2.5 mM).

Cycling Conditions Single PCR

Cycling Conditions Multiplex PCR

1× 2′/ 95°C 35× (1′/ 95°C, 1′/ 62°C, 1′/72°C) 1× 5′/72°C 1× 4′/ 94°C 35× (3′0′/ 94°C, 30′′/ 55°C, 30′′/72°C) 1× 5′/72°C 1× 2′/ 95°C 35x (1′/ 95°C, 1′/ 62°C, 1′/72°C) 1× 5′/ 72°C

Amplicons (bp)

809

1× 2′/ 95°C 35× (1’/95°C, 1′/ 62°C, 1′/72°C) 1× 5′/ 72°C

260

372

540

Molecular Detection of Foodborne Pathogens

(2) Perform PCR amplification using the cycling conditions outlined below, and separate by 1.5% agarose gel containing 0.5 μg/ml ethidium bromide.117 (ii) Quantitative PCR for A. westerdijkiae:

80

(1) Prepare PCR mixture (50 μl) containing Platinum® SYBR® Green qPCR Supermix-UDG (Invitrogen Life Technologies), Bt2Aw-F (0.4 μM)/Bt2Aw-R (0.4 μM) primers, and 2.0 μl of total DNA.

Species A. westerdijkiae

Primers (5′–3′) Bt2Aw-F TGATACCTTGGCGCTTGTGACG Bt2Aw-R CGGAAGCCTAAAAAATGAAGAG

(2) Perform PCR amplification using the cycling conditions outlined above. Note: For A. westerdijkiae DNA detection in coffee beans, 2.0 μl of each extract diluted 100-fold is added to the reaction mixture to record the cycle threshold (Ct) value. Quantification of the unknown samples is carried out by interpolating the samples Ct in a standard curve. To construct the standard curve a fragment of the Btub gene (600 bp) is first amplified, by using Bt2a and Bt2b primers.118 After purification, the amount of PCR product is determined in a fluorometer (DyNa Quant 200; Pharmacia, Uppsala, Sweden). DNA amounts are converted in gene copy number by using the formula mentioned below:

Species

A. carbonarius

Primers (5′–3′) CARBO_q1 CCGATGGAGGTCATGACATGA CARBO_q2 AATGCGAACCGGATATTAACTTCTG CARBO_probe 6FAM-CAGCGGCGGAGATA-MGB

Cn (copy number/ml)    Fragment concentration (mg/l) =  1.004 × 1015 Fragment size (bp) The quantified amplicon is ten-fold serially diluted (109–101 Cn) and then used as template DNA in a real-time reaction using Bt2Aw-F and Bt2Aw-R as primers. The specificity is checked through melting curve analysis. The sensitivity of

this real-time PCR method is less than ten haploid genomes per 0.1 g of coffee beans. 38.2.1.2 Grapes Sample preparation. With a clean spatula, excise a small amount (about 200 mg) of the grinded sample and transfer it into the labeled 1.5-ml microcentrifuge tube and add 400 µl Buffer SP1 supplied with the MagBead Plant DNA Kit (Machercy-Nayel) and 2 µl RNase A. Follow kit protocol steps until step 5; Carefully transfer 200 µl of cleared

Cycling Conditions Single PCR

Amplicons (bp)

1× 2′/50°C 1× 5′/95°C 45× (30′′/95°C, 30′′/ 62°C, 30′′/72°C)

347

supernatant to a 1.5-ml centrifuge tube and add 4 µl of Exo IPC DNA (50×) (Applied Biosystems). Then continue steps according to the kit protocol. Detection procedure. Quantitative PCR for A. ­carbonarius 84 (1) Prepare PCR mixture (25 µl) containing 12.5 µl of 2× Taqman PCR Master Mix (Applied Biosystems), 0.9 µl of a master mix: CARBO_q1 (50 nM), CARBO_q2 (300 nM) and CARBO_probe (75 nM), 2.5 µl 10× Exo IPC Mix (IPC primers and Vicprobe), 5 µl DNA grapes sample diluted 1:10 (food sample assay) or dilutions of A. carbonarius DNA (fungal DNA reference curve) or dilutions of Exo

Cycling Conditions Single PCR

Amplicons (bp)

1× 2′/50°C 1× 10′/95°C 45× (15′′/ 95°C, 1′/60°C)

168

Target Gene

Calmodulin

IPC DNA (Control DNA reference curve), and 4.1 µl double-distilled H2O (2) Mix reaction reagents making a reaction cocktail containing all of the common reagents. Change pipette tips between all stocks of reagents to avoid cross-contamination of the different tubes of solutions. Mix each reaction cocktail in a separate labeled 1.5-ml microcentrifuge tube. Keep the

541

Aspergillus

reaction mixes on ice until dispensed into the final reactions. (3) Perform PCR amplification in triplicates for each grapes sample and standard using the conditions outlined in ABI PRISMR 7000 (Applied Biosystems, Foster City, CA) or similar apparatus for qPCR amplification and detection.

Species

A. flavus A. parasiticus

Genes

Amplicons (bp)

aflD

400

aflM

537

aflP

797

Detection procedure. Qualitative PCR for A. flavus and A. parasiticus using target genes aflD aflM aflP97 (1) Prepare PCR mixture (50 µl) containing 5 µl template DNA (2 ng/ml), 1.25 µl primer (120 pmol/ml each), 200 µM dNTPs, Taq polymerase buffer 10× and 1 U Taq polymerase.

Primers (5′–3′) nor1 accgctacgccggcactctcggcac nor2 gttggccgccagcttcgacactccg ver1 gccgcaggccgcggagaaagtggt ver2 ggggatataccccgcgacacagcc omt-1 gtggacggacctagtccgacatcac omt-2 gtcggcgccacgcactgggttgggg

Note: Calculate standard curves for quantification purposes using: (i) ten-fold serial dilutions of purified DNA from A. carbonarius (ITEM 4780), ranging from 5×101–5×10 −4 ng (determined by a spectrophotometer), to evaluate A. carbonarius presence; (ii) exogenous IPC DNA (10×, 1×, 0.1×), to evaluate the yield of DNA isolation step for each sample and to correct quantitative results for A. carbonarius DNA in grapes samples for relative recovery rate. Standard curves generated plotting different DNA amounts against relatives Ct values allow to interpolate Ct values (cycle number at which the fluorescence generated within a reaction crosses the threshold line, rising above the base line and becoming statistically significant) obtained for unknown samples analyzed.

38.2.2 Molecular Detection of Aflatoxigenic Aspergillus Species in Food 38.2.2.1  Figs Sample preparation. (1) Separate the surface area of the fig tissue and freeze it in liquid nitrogen. (2) To extract DNA grind the material in a mortar and resuspend in lysis buffer. (3) Incubate the suspension at 68°C for 15 min, then centrifuge for 15 min at 15000×g. (4) After centrifugation, transfer a volume of 7 ml of the supernatant to a new centrifuge tube and add 1 ml of 4 M sodium acetate. (5) Place the solution on ice for 1 h and centrifuge for 15 min at 15000×g. (6) After centrifugation, transfer 6 ml of the supernatant to a fresh tube. (7) Extract solution with phenol and precipitate the isolated DNA by the addition of 2.5 volumes of ethanol.

Cycling Conditions

30× (1′/ 95°C, 2′/65°C, 4′/72°C)

(2) Perform PCR amplification using the cycling conditions outlined above. (3) Analyze the PCR products by electrophoresis on agarose gel stained with ethidium bromide. Note: The minimal detection concentration with purified fungal DNA is 25 pg DNA per reaction; when DNA from figs is present in the reaction mixture, the sensitivity of the reaction decreases by a factor of 10 (250 pg per reaction). When DNA isolated from infected figs is present in the reaction with the aflP specific primer set, two additional bands appear and the PCR product specific for the aflM gene is very weak. 38.2.2.2  Corn Sample preparation. (1) Pulverize 1 g of plant material with a mortar and pestle. (2) Add 7 ml of lysis buffer and mix gently. (3) Add 50 µl of RNase cocktail and incubate the mixture at 37°C for 30 min. (4) Add 1 ml of Proteinase K and incubate at 50°C for 1 h. (5) Perform a double phenol-chloroform extraction followed by isopropanol precipitation. (6) Resuspend in 200 µl of TE buffer. Detection procedure. Qualitative PCR for A. flavus and A. parasiticus using target genes aflR aflM aflP102 (1) Prepare PCR mixture (50 µl) containing 100 ng template DNA, 1 µM each primers, 200 µM dNTPs, Taq polymerase buffer 10×, and 2.5 U Taq polymerase. (2) Perform PCR amplification using the cycling conditions outlined above.

542

Species

A. flavus A. parasiticus

Molecular Detection of Foodborne Pathogens

Genes

Amplicons (bp)

aflR

1032

aflM

895

aflP

1024

Primers (5′–3′) APA450 tatctccccccgggcatctcccgg APA1482 ccgtcagacagccactggacacgg VER496 atgtcggataatcaccgtttagatggc VER1391 cgaaaagcgccaccatccaccccaatg OMT208 ggcccggttccttggctcctaagc OMT1232 cgccccagtgagacccttcctcg

Cycling Conditions 30/40× (1′/94°C, 2′/65°C, 2′/72°C)

(3) Analyze the PCR products by electrophoresis on agarose gel stained with ethidium bromide.

Detection procedure. Qualitative multiplex PCR for A. parasiticus using target genes aflR aflD aflM aflP107

Note: Regardless of the primer set used, amplification patterns are obtained from A. parasiticus DNA. When A. flavus DNA is present, 40 PCR cycles are necessary to see a low intensity band for aflM gene.

(1) Prepare PCR mixture (20 µl) containing 2 µl DNA template, 8 µl of the premixed primer solution (containing 5 pmol of each of eight primers), 4 µl of 250 µM dNTP, Taq polymerase buffer 10× and 2 U Taq DNA polymerase.

Species

A. parasiticus

Gene

Amplicons (bp)

Primers (5′-3′)

aflR

1032

aflD

400

aflM

538

aflP

1025

APA450 tatctccccccgggcatctcccgg APA1482 ccgtcagacagccactggacacgg nor1 accgctacgccggcactctcggcac nor2 gttggccgccagcttcgacactccg ver1 gccgcaggccgcggagaaagtggt ver2 ggggatataccccgcgacacagcc omt-1 gtggacggacctagtccgacatcac omt-2 gtcggcgccacgcactgggttgggg

38.2.2.3  Peanut Kernel Sample preparation. (1) Seal freshly harvested peanut kernel in polypropylene plastic bags and store at –25°C until used. (2) For genomic DNA extraction from an individual kernel placed in a bag, remove the heart and break a cotyledon into pieces by hands. (3) Take a cotyledon piece about the same size as the germ and weigh. (4) Extract genomic DNA from the germ and the cotyledon (ca. 20 mg each) using a commercial DNA extraction kit. (DNeasy Plant Mini Kit, Piager) (5) Grind the samples with 400 µl of kit lysis buffer in a mortar with liquid nitrogen. (6) After grinding, remove the suspension with a disposable pipette to a 1.5 ml microfuge tube. (7) Follow the recommended procedure for DNA extraction.

Cycling Conditions

35× (1′/94°C, 2′/65°C, 2′/72°C)

(2) Perform PCR amplification using the cycling conditions outlined above. (3) After reaction, load 10 µl of the PCR product onto agarose gel and visualise after staining with ethidium bromide. Note: The target DNA fragments are detected in the kernels infected with A. parasiticus, and none is detected in the sound kernels or in the kernels infected with A. niger. 38.2.2.4 Wheat Sample preparation. (1) Freeze 0.5 g of the infected sample in liquid nitrogen and grind to powder in a mortar. (2) Use an amount of 200 mg of the powder for isolation of total RNA. Use the recommendations of the manufacturer of the kit (E.Z.N.A fungal RNA kit, Peqlab) for RNA extraction. (3) Treat the RNA preparation with DNase I for degradation of traces of genomic DNA.

543

Aspergillus

(4) Check the integrity of the RNA on agarose gel. (5) Use the DNase I-treated total RNA for cDNA synthesis which will be used for real-time PCR or stored at –20°C. Detection procedure. Quantitative real-time reverse transcription PCR for A. flavus using target gene aflR110 (1) Prepare PCR mixture (50 µl) containing 1 µl DNA sample, 0.5 µl of the primers and probe (nortaq-1, nortaq-1 and norprobe, each 0.5 µM), 1 µl of each dNTP, 7 µl of 25 mM MgCl2, TaqMan buffer A 10×, 0.5 µl of uracil-N-glycosylase, and 1 U Taq DNA polymerase.

Species

Gene

applied to the screening of agricultural commodities for the presence of mycotoxin producers prior to or even after processing, and negative results may indicate that the sample is virtually free of mycotoxins. Otherwise, the positive samples must be analyzed for the presence of mycotoxins, although it seems to be even possible to correlate the number of target molecules with fungal biomass and indirectly with mycotoxins content by using a quantitative approach.84,85 Most of the developed methods have been proven to be robust and reliable and are ready to be applied at the practical level. Their sensitivity is high and often exceeds that of comparable methods, so conventional PCR could become a quick routine method to check if some mycotoxins producing Aspergillus fungi are present in a food sample, and

Primer (5′–3′)

Cycling Conditions

nortaq-1 gtccaagcaacaggccaagt nortaq-2 tcgtgcatgttggtgatggt norprobe tgtcttgatcggcgcccg A. flavus

aflR nor1 accgctacgccggcactctcggcac nor2 gttggccgccagcttcgacactccg bentaq1 cttgttgaccaggttgtcgat bentaq2 gtcgcagccctcagcct benprobe cgatgttgtccgtcgcgaggct

1× 2′/50°C 1× 10′/95°C 35× 20′′/94°C, 20′′/65°C, 30′′/72°C

β−tub ben1 ggccagtccggtgctcc ben2 gttgtcgatacagaagg

(2) Perform PCR amplification using conditions outlined above. Note: Generate the standard curve using a larger PCR fragment of the aflR gene amplified with the primers nor-1/nor-2. Determine the concentration of this standard PCR product in a fluorometer and calculate the number of copies. The stock solutions are diluted serially by a factor of 10, and an aliquot of the dilutions is used as copy number standard. The same reaction conditions are used with the β-tubulin gene (ben A56) as the target sequence. AflR mRNA can be detected 48 h before the first AFB1 could be determined

38.3 Conclusions and Future Perspectives This chapter has illustrated in various traits the potential of PCR systems to differentiate between morphologically similar species having different toxicological profiles, thus these methods are useful tools for the objective assessment of food safety by identifying mycotoxins producing Aspergillus species that are difficult to characterize. PCR methods may be

to evaluate the toxicological potential risks. However, the choice of unique target sequences is vital for developing molecular detection methods. For the mycotoxigenic fungi it is preferred to base these methods on single or multiple genes of biosynthetic pathways as described for aflatoxigenic Aspergillus; but in the case of biosynthetic genes not yet elucidated, such as the ochratoxigenic genes, anonymous markers or unique conserved gene sequences (ITS, 18S, calmodulin, beta-tubulin, etc.) must be chosen. As gene/s of biosynthetic pathways used as targets could be more promising in correlation to mycotoxins contamination and are able to amplify one or more producing species in a unique PCR reaction; primers developed on anonymous marker or gene are highly species-specific and detect the potential producers without discriminating between producing and nonproducing strains. In this case the molecular system needs more than one primer possibly in a multiplex reaction for detecting a group of AFs or OTA producing Aspergillus. The trial to differentiate among producing and nonproducing strains by means of DNA PCR-methods are based on the assumption that a genetic difference, which is responsible for the mycotoxins negative phenotype, can be detected by PCR

544

using biosynthetic genes. Since these genes are supposed to be exclusively present in organisms potentially producing mycotoxins, their use as targets may increase accuracy, sensitivity and specificity of detection assays. However, positive or negative results may not be directly correlated to mycotoxin production: mutations or variations could not be located in the target sequence, or, if they are present, the gene functionality could not be compromised. For example some genes needed for aflatoxin biosynthesis such as aflD, aflM, aflP, and aflR are present in the ­nonaflatoxin producing fungi A. oryzae and A. sojae, although they are not expressed. In addition, false positives may frequently arise since the technique cannot detect mutation outside the primers’ targeted region of gene sequence. Numerous studies have documented that different parameters such as water activity, pH, temperature, nutritional factors, and even the concomitant presence of other microrganisms, have a significant influence on the growth of fungal cells and thereby on aflatoxin production. The mycotoxin biosynthetic genes are not expressed constitutively, therefore their activation, under suitable conditions, can be used to monitor the mycotoxin biosynthesis. The monitoring of the activity of biosynthetic genes, by measuring their expression levels, appears as a better way to distinguish between toxigenic and nontoxigenic strains. This method is closer to the phenotypic “front” than the DNA based approach, as it gives information about the expression of the genes. In this regard, three expression monitoring systems based either on conventional RT-PCR112,114 or RT real-time PCR110 have been developed for aflatoxigeni fungi. Mayer demonstrated the correlation between induction of the gene and production of AFs even 48 h before the detection of the mycotoxin in the substrate. So, the molecular methods have proved successful in measuring the process of induction (gene activation) in the formation of mycotoxins on the basis of environmental parameters. Then the combination of this information allowing the activation of mycotoxins genes can be regarded as molecular critical control point (MCCP’s), and the capacity of this molecular system to detect the induction of toxin genes before the production occurs can be used as online monitoring systems.64 This is mainly the case of PCR diagnostic methods for aflatoxigenic fungi developed for A. flavus and A. parasiticus, which are the major species of concern for production of aflatoxin in agricultural commodities and share a 90–99% identity in the aflatoxin clustered genes.4 A PCR detection system based on biosynthetic genes has not yet been described for the rarely encountered aflatoxin producers A. nomius, A. bombycis, A. ochraceoroseus, and A. pseudotamari, but the presence of homologues of the biosynthetic genes may be hypothesized also in these minor aflatoxin producing species and so their use as targets in detection PCR assays. Another important point is that, despite there being four main aflatoxins naturally produced by Aspergilli, no PCR methods have been developed to differentiate the specific compounds, aflatoxin B1 being the most toxic and so the principal target. An interesting issue is raised also by the shared metabolic pathway for aflatoxins and sterigmatocystin, toxic intermediate of the

Molecular Detection of Foodborne Pathogens

aflatoxin biosynthesis produced mainly by A. versicolor and A. nidulans. Therefore primers should be tested with sterigmatocystin producers to avoid false-positive results in the monitoring of aflatoxin producers. As the most advanced PCR systems are based on mycotoxins biosynthetic genes, an understanding and exploration of the OTA biosynthetic genes pathway in Aspergillus will give a great advantage in the future PCR systems for monitoring OTA producing Aspergillus species and correlate them to food contamination by OTA. However, the available assays, which detect potential producers of OTA, have been shown to work well with pure fungal cultures, but applicability and usefulness in the routine analysis of commodities remains to be demonstrated for most of them. Therefore, capability of detection their DNA target organisms in contaminated sample materials is an essential step if assays are to be used in practical applications. Finally, it is necessary and fundamental to further develop PCR-based assays on foodborne Aspergillus species for their application in quality food control and hazard analysis applications. In this respect, false-negative reactions must be distinguishable from true negative reactions (use of internal DNA control); extensive testing for specificity of target sequences, optimization of sensitivity in sample materials, rapidity and ease of use for trained and untrained personnel are also necessary. Also sampling, sample preparation and DNA extraction must further be optimized and tested with high numbers of samples. In this context the validation of PCR methods for food safety related to fungi and mycotoxins contamination risk must be performed throughout an interlaboratory study involving independent laboratories which all analyze the same samples under standardized conditions. With the aim of standardization of methods and protocols, various national and international agencies are currently defining the conditions that must be accomplished to routinely use PCR in assessing safety of food samples. For example, primers targeting the aflatoxin biosynthetic genes in the various PCR assays, have been usually chosen among those already used by different research groups, allowing comparison among results. To date, the standardization of PCR protocols in food has been defined and described for bacteria, but none have been outlined for PCR detection of fungi in food. PCR methods are useful in monitoring quality and safety of food; successively the quantitative approach of real-time PCR has been applied to correlate either the DNA content to fungal biomass or even mycotoxins content, or to quantify mRNA in monitoring gene expression involved in mycotoxins biosynthesis under different conditions and food matrices. The most promising breakthroughs in this field are expected to be in the area of sensor technology, as bio-sensors can be used to detect different targets like pathogens, pesticides, and toxins. Among the different types of biosensors, the microarray technology has been widely studied and applied in the last five years and has increased the diagnostic efficacy of the molecular methods, e.g., in monitoring the expression profile of all the biosynthetic cluster

545

Aspergillus

genes of a particular mycotoxin. Further development for toxigenic fungi detection might include microarray DNA or RNA chips. With the capacities available in microarray technology, probes for biosynthetic genes involved in the production of different relevant mycotoxins, e.g., aflatoxins, trichothecenes, fumonisins, OTA, might be combined aimed at optimizing food production processes for minimized risk of mycotoxin production. Detailed expression profiles of whole mycotoxin biosynthetic pathways or groups of key genes can be generated by RNA microarray allowing the exact prediction and control of mycological status of foods. With this aim, Schmidt-Heydt and Geisen119 developed a microarray carrying oligonucleotides for the biosynthetic pathways of the most known and relevant mycotoxins (aflatoxins, ochratoxin, fumonisins, trichothecene, and patulin). The microarray was highly specific in showing the differences in mycotoxins pathway gene expression on various media and growth conditions. Also, it is designed in a way allowing to add newly mycotoxins pathway genes, and it demonstrates its functionality on natural food commodities like wheat. Molecular detection systems such as microarray could be used and developed in future to systematically either investigate the influence of relevant parameters in mycotoxins gene expression, or monitor the quality and safety of food products.







References



1. Powell, K.A., Renwick, A. and Peberdy, J.F. The Genus Aspergillus: From Taxonomy And Genetics To Industrial Applications. Plenum Press, New York, 1994. 2. Smith, J.E. Aspergillus, Biotechnology Handbooks, Vol. 7. Plenum Press, New York, 1994. 3. Lam, T.Y. US Patent 4, 376, 863, 1983. 4. Bennett, J.W. and Klich, M.A Aspergillus: Biology and Industrial Applications. Butterworth-Heinemann, Boston, MA, 1992. 5. Raper, K.B. and Fennel, D.I. The Genus Aspergillus. William & Wilkins, Baltimore, MD, 1965. 6. Varga, J. et al. Molecular diversity of agriculturally important Aspergillus species. Eur. J. Plant Pathol., 110, 627, 2004. 7. Bennett, J.W. and Klich, M. Mycotoxins. Clin. Microbiol. Rev., 16, 497, 2003. 8. Payne, G.A. and Brown, M.P. Genetics and physiology of aflatoxin biosynthesis. Annu. Rev. Phytopathol., 36, 329, 1998. 9. Yu, J. et al. Clustered pathway genes in aflatoxin biosynthesis. Appl. Environ. Microbiol., 70, 1253, 2004. 10. Krishnamachari, K.A. et al. Hepatitis due to aflatoxicosis. An outbreak in werstern India. Lancet, 1, 1061, 1975. 11. Azziz-Baumgartner, E. et al. Case-control study of an acute aflatoxicosis outbreak, Kenya 2004. Environ. Health Perspect., 113, 1779, 2005. 12. IARC Monogr. Eval. Carcinog. Risks. Hum. Some naturally occurring substances: food items and constituents, heterocyclic aromatic amines and mycotoxins. International Agency for Research on Cancer, Lyon, France, 56, 489, 1993. 13. EFSA (European Food Safety Authority). Opinion of the Scientific Panel on contaminants in the Food Chain of the EFSA on a request from the Commission related to ochratoxin A in food. EFSA J., 365, 1, 2006. Available from:http://





www.efsa.europa.eu/etc/medialib/efsa/science/contam/ contam_­opinions/1521.Par.0001.File.dat/contam_op_ej365_ ochratoxin_a_food_en1.pdf 14. Petzinger, E. and Weidenbach, A. Mycotoxins in the food chain: the role of ochratoxins. Livest. Prod. Sci., 76, 245, 2002. 15. Samson, R.A. et al. New ochratoxin A or sclerotium producing species in Aspergillus section Nigri. Stud. Mycol., 50, 45, 2004. 16. Frisvad, J.C. et al. New ochratoxin A producing species of Aspergillus section Circumdati. Stud. Mycol., 50, 23, 2004. 17. Peterson, S. W. et al. Phylogeny and subgeneric taxonomy of Aspergillus. In Aspergillus in the Genomic Era, p. 33, Varga, J. and Samson, R.A. (Eds.). Wageningen Academic Publishers, Wageningen, The Netherlands, 2008. 18. Gams, W. et al. Infrageneric taxa of Aspergillus. In Advances in Aspergillus and Penicillium Systematics, p. 55, Samson, R.A. and Pitt, J.I., (Eds.). Plenum Press, New York, 1985. 19. Pitt, J.I. and Hocking, A.D. Fungi and Food Spoilage. Blackie Academic and Professional, London, UK, 1997. 20. Klich, M.A. Identification of Common Aspergillus Species. Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands, 2002. 21. Samson, R.A, Hoekstra, E.S. and Frisvad, J.C. Introduction to Food- and Air-borne Fungi, 7th edition. Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands, 2004. 22. Peterson, S.W., Phylogenetic relationships in Aspergillus based upon rDNA sequences analysis. In Integration of Modern Taxonomic Methods for Penicillium and Aspergillus Classification, Samson, R.A. and Pitt, J.I. (Eds.). Harwood Academic Publishers, Amsterdam, 2000. 23. Rigó, K. et al. Phylogenetic analysis of Aspergillus section Flavi based on sequences of the internal transcribed spacer regions and the 5.8 rRNA gene. J. Gen. Appl. Microbiol., 48, 9, 2002. 24. Samson, R.A. A compilation of the Aspergilli described since 1965. Stud. Mycol., 18, 1, 1979. 25. Frisvad, J.C., Skouboe, P. and Samson, R.A. Taxonomic comparison of three different groups of aflatoxin producers and a new efficient producer of aflatoxin B1, sterigmatocystin and 3-O-methylsterigmatocystin, Aspergillus rambellii sp. nov. Syst. Appl. Microbiol., 28, 442, 2005. 26. Pildain, M.B. Two novel aflatoxin-producing Aspergillus species from Argentinean peanuts. Int. J. Syst. Evol. Microbiol., 58, 725, 2008. 27. Samson, R.A., Hong, S.B. and Frisvad, J.C. Old and new concepts of species differentiation in Aspergillus. Med. Mycol., 44, S133, 2006. 28. Samson, R.A. et al. Diagnostic tools to identify black aspergilli. Stud. Mycol., 59, 129, 2007. 29. Perrone, G. et al. Biodiversity of Aspergillus species in some important agricultural products. Stud. Mycol., 59, 53, 2007. 30. Serra, R. et al. Aspergillus ibericus: a new species of the section Nigri isolated from grapes. Mycology, 98, 295, 2006. 31. Perrone, G. et al. Aspergillus uvarum sp. nov., a uniseriate black Aspergillus species isolated from grapes in Europe. Int. J. Syst. Evol. Microbiol., 58, 1032, 2008. 32. Battilani, P., Giorni, P. and Pietri, A. Epidemiology of toxin producing fungi and ochratoxin A occurrence in grape. Eur. J. Plant Pathol., 109, 715, 2003. 33. Leong, S.L. et al. Australian research on ochratoxigenic fungi and ochratoxin A. Int. J. Food Microbiol., 111 (Suppl. 1), S10, 2006. 34. Medina, A. et al. Study of Spanish grape mycobiota and ochratoxin A production by isolates of Aspergillus tubingensis and other members of Aspergillus section Nigri. Appl. Environ. Microbiol., 71, 4696, 2005.

546 35. Perrone, G. et al. Ochratoxin A production and AFLP analysis of Aspergillus carbonarius, Aspergillus tubingensis, and Aspergillus niger strains isolated from grapes in Italy. Appl. Environ. Microbiol., 72, 680, 2006. 36. Dalcero, A. et al. Detection of ochratoxin A in animal feeds and capacity to produce this mycotoxin by Aspergillus section Nigri in Argentina. Food Addit. Contam., 19, 1065, 2002. 37. Ponsone, M.L. et al. Ochratoxin A and ochratoxigenic Aspergillus species in Argentinian wine trapes cultivated under organic and non-organic systems. Int. J. Food Microbiol., 114, 131, 2007. 38. Bayman, P. et al. Ochratoxin production by the Aspergillus ochraceus group and Aspergillus alliaceus. Appl. Environ. Microbiol., 68, 2326, 2002. 39. Perrone, G. et al. Aspergillus in grapes: ecology, biodiversity and genomics. In Aspergillus in the Genomic Era, Varga, J. and Samson, R.A. (Eds.). Wageningen Academic Publishers, Wageningen, The Netherlands, 2008. 40. Pardo, E. et al. Ecophysiology of ochratoxigenic Aspergillus ochraceus and Penicillium verrucosum isolates. Predictive models for fungal spoilage prevention—a review. Food Addit. Contam., 23, 398, 2006. 41. Esteban, A. et al. Effects of temperature and incubation time on production of ochratoxin A by black Aspergilli. Res. Microbiol., 155, 861, 2004. 42. Joosten, H.M.L.J. et al. Production of ochratoxin A by Aspergillus carbonarius on coffe cherries. Int. J. Food Microbiol., 65, 39, 2001. 43. Taniwaki, M.H. et al. The source of ochratoxin A in Brazilian coffee and its formation in relation to processing methods. Int. J. Food Microbiol., 82, 173, 2003. 44. Belli, N. et al. Aspergillus carbonarius growth and ochratoxin A production on a synthetic grape medium in relation to environmental factors. J. Appl. Microbiol., 98, 839, 2005. 45. Mitchell, D., Aldred, D. and Magan, N. Impact of ecological factors on the growth and ochratoxin A production by Aspergillus carbonarius from different regions of Europe. Asp. Appl. Biol., 68, 109, 2003. 46. Bayman, P. and Baker, J.L. Ochratoxins: a global perspective. Mycopathologia, 162, 215, 2006. 47. Klich, M. Aspergillus flavus: the major producer of aflatoxin. Mol. Plant Pathol., 8, 713, 2007. 48. Sanchis, V. and Magan, N. Environmental conditions affecting mycotoxins. In Mycotoxins in Food, Magan, N. and Olsen, M. (Eds.). Woodhead Publishing Limited, Cambridge, UK, 2004. 49. Hedayati, M.T. et al. Aspergillus flavus: human pathogen, allergen and mycotoxin producer. Microbiology, 153, 1677, 2007. 50. Klich, M. Environmental and developmental factors influencing aflatoxin production by Aspergillus flavus and Aspergillus parasiticus. Mycoscience, 48, 71, 2007. 51. Horn, W.B. Biodiversity of Aspergillus section Flavi in the United States: a review. Food Addit. Contam., 24, 1088, 2007. 52. Klich, M.A. et al. Phylogenetic and morphological analysis of Aspergillus ochraceoroseus. Mycologia, 95, 1252, 2003. 53. Geiser, D.M. et al. The phylogenetic of mycotoxin and sclerotium production in Aspergillus flavus and Aspergillus oryzae. Fungal Genet. Biol., 31, 169, 2000. 54. Geiser, D.M. et al. Cryptic speciation and recombination in the aflatoxin producing fungus Aspergillus flavus. Proc. Natl. Acad. Sci. USA, 95, 388, 1998. 55. Pitt, J.I., Hocking, A.D. and Glenn, D.R. An improved medium for the detection of Aspergillus flavus and A. parasiticus. J. Appl. Bacteriol., 54, 109, 1983.

Molecular Detection of Foodborne Pathogens 56. Heenan, C.N., Shaw, K.J. and Pitt, J.I. Ochratoxin production by Aspergillus carbonarius and A. niger isolates and detection using coconut cream agar. J. Food Mycol., 1, 67, 1998. 57. Rojas-Durán, T.R. et al. Study of a room temperature phosphorescence phenomenon to allow the detection of aflatoxigenic strain in culture media. Int. J. Food Microbiol., 115, 149, 2007. 58. Zhao, J. et al. Identification of Aspergillus fumigatus and related species by nested PCR targeting ribosomal DNA internal transcribed spacer regions. J. Clin. Microbiol., 39, 2261, 2001. 59. Varga, J. et al. Aspergillus brasiliensis sp. nov., a biseriate black Aspergillus species with world wide distribution. Int. J. Syst. Evol. Microbiol., 57, 1925, 2007. 60. Pringle., A. et al. Cryptic speciation in the cosmopolitan and clonal human pathogenic fungus Aspergillus fumigatus. Evolution Int. J. Org. Evol., 59, 1886, 2005. 61. Balajee, S.A. et al. Aspergillus lentulus sp. nov., a new sibling species of A. fumigatus. Eukaryot. Cell., 4, 625, 2005. 62. Geiser, D.M. Practical molecular taxonomy of fungi. In Advances in Fungal Biotechnology for Industry, Medicine and Agriculture, Lange, L. and Tkacz, J. (Eds.). Kluwer Academic Publishers, New York, 2004. 63. Geiser, D.M. et al. The current status of species recognition and identification in Aspergillus. Stud. Mycol., 59, 1, 2007. 64. Geisen, R. Molecular detection and monitoring of fungi. In Food Mycology: A Multifaced Approach to Fungi and Food, Mycology Series Vol. 25. Dijksterhuis, J. and Samson, R.A. (Eds.), CRC Press, Boca Raton, Florida, 2007. 65. Varga, J. Molecular typing of aspergilli: recent developments and outcomes. Med. Mycol. 44, 149, 2006. 66. Bain, J.M. et al. Multilocus sequence typing of the pathogenic fungus Aspergillus fumigatus. J. Clin. Microbiol., 45, 1469, 2007. 67. de Ruiter, M.T. et al. Retrotransposon insertion-site context (RISC) typing: a novel typing method for Aspergillus fumigatus and a convenient PCR alternative to restriction fragment length polymorphism analysis. J. Microbiol. Methods, 70, 528, 2007. 68. O’Callaghan, J., Caddick, M.X., and Dobson, A.D.W. A polyketide synthase gene required for ochratoxin A biosynthesis in Aspergillus ochraceus. Microbiology, 149, 3485, 2003. 69. O’Callaghan, J., Stapleton, P.C. and Dobson, A.D.W. Ochratoxin A biosynthetic genes in Aspergillus ochraceus are differentially regulated by pH and nutritional stimuli. Fungal Genet. Biol., 43, 213, 2006. 70. Niessen, L. Current trends in molecular diagnosis of ochratoxin A producing fungi. In Mycotechnology—Present Status and Future Prospects, Rai, M. (Ed.). I.K. International Publishing House, New Delhi, India, 2006 . 71. Schmidt, H. et al. Molecular typing of Aspergillus ochraceus and construction of species specific SCAR-primers based on AFLP. Syst. Appl. Microbiol., 26, 138, 2003. 72. Schmidt, H. et al. Utilization of AFLP markers for PCR— based identification of Aspergillus carbonarius and indication of its presence in green coffee samples. J. Appl. Microbiol., 97, 899, 2004. 73. Fungaro, M.H.P. et al. A molecular method for detection of Aspergillus carbonarius in coffee beans. Curr. Microbiol., 49, 123, 2004. 74. Jimenez, L. et al. Molecular detection and identification of Aspergillus niger contamination in cosmetic/pharmaceutical raw materials and finished products. J. Rapid Methods Aut. Microbiol., 7, 39, 1999.

Aspergillus 75. Sugita, T. et al. PCR identification system for the genus Aspergillus and three major pathogenic species: Aspergillus fumigatus, Aspergillus flavus and Aspergillus niger. Med. Mycol., 42, 433, 2004. 76. Perrone, G. et al. PCR assay for identification of Aspergillus carbonarius and Aspergillus japonicus. Eur. J. Plant Pathol., 110, 641, 2004. 77. Gonzàlez-Salgado, A. et al. Discrimination of Aspergillus niger and other Aspergillus species belonging to section Nigri by PCR assays. FEMS Microbiol. Lett., 245, 353, 2005. 78. Susca, A., et al. Polymerase chain reaction (PCR) identification of Aspergillus niger and Aspergillus tubingensis based on the calmodulin gene. Food Addit. Contam., 24, 1154, 2007. 79. Patino, B. et al. PCR detection assays for the ochratoxin­producing Aspergillus carbonarius and Aspergillus ochraceus species. Int. J. Food Microbiol., 104, 207, 2005. 80. Morello, L.G. et al. Detection and quantification of Aspergillus westerdijkiae in coffee beans based on selective amplification of β-tubulin gene by using real-time PCR. Int. J. Food Microbiol., 119, 270, 2007. 81. Susca, A., Stea, G. and Perrone, G. Rapid polymerase chain reaction (PCR)-single-stranded conformational polymorphism (SSCP) screening method for the identification of Aspergillus section Nigri species by the detection of the calmodulin nucleotide variations. Food Addit. Contam., 24, 1148, 2007. 82. Bufflier, E. et al. detection of Aspergillus carbonarius and other black aspergilli from grapes by DNA OLISATM microarray. Food Addit. Contam., 24, 1138, 2007. 83. Schaad, N.W. and Frederick, R.D. Real-time PCR and its application for rapid plant disease diagnostics. Can. J. Plant Pathol., 24, 250, 2002. 84. Mulè, G. et al. Development of a quantitative real-time PCR assay for the detection of Aspergillus carbonarius in grapes. Int. J. Food Microbiol., 111, S28, 2006. 85. Atoui, A., Mathieu, F. and Lebrihi, A. Targeting a polyketide synthase gene for Aspergillus carbonarius quantification and ochratoxin A assessment in grapes using real-time PCR. Int. J. Food Microbiol., 115, 313, 2007. 86. Selma, M.V, Martinez-Culebras, P.V and Aznar, R. Realtime PCR based procedures for detection and quantification of Aspergillus carbonarius in wine grapes. Int. J. Food Microbiol., 122, 126, 2008 87. Schmidt, H. et al. Detection and quantification of Aspergillus ochraceus in green coffee by PCR. Lett. Appl. Microbiol., 38, 464, 2004. 88. Seifert, K.A. and Lévesque, C.A. Phylogeny and molecular diagnosis of mycotoxigenic fungi. Eur. J. Plant Pathol., 110, 449, 2004. 89. Geisen, R. et al. Development of a real time PCR system for detection of Penicillium nordicum and for monitoring ochratoxin A production in foods by targeting the ochratoxin polyketide synthase gene. Syst. Appl. Microbiol., 27, 501, 2004. 90. Geisen, R. Schmidt-Heydt, M. and Karolewiez, A., A gene cluster of the ochratoxin A biosynthetic genes in Penicillium. Mycot. Res., 22, 134, 2006. 91. Atoui, A. et al. Amplification and diversity analysis of ketosynthase domains of putative polyketide synthase genes in Aspergillus ochraceus and Aspergillus carbonarius producers of ochratoxin A. Mol. Nutr. Food Res., 50, 448, 2006. 92. Dao, H.P., Mathieu, F. and Lebrihi, A. Two primer pairs to detect OTA producers by PCR method, Int. J. Food Microbiol., 104, 61, 2005.

547 93. Vrabcheva, T. et al. Cooccurrence of ochratoxin A and citrinin in cereals from Bulgarian villages with a history of Balkan endemic nephropathy. J. Agric. Food Chem., 48, 2483, 2000. 94. Yu, J., Bhatnagar, D. and Ehrlich, K.C. Aflatoxin biosynthesis. Rev. Ibeoram. Micol., 19, 191, 2002. 95. Bhatnagar, D., Ehrlich, K.C. and Cleveland, T. E. Molecular genetic analysis and regulation of aflatoxin biosynthesis. Appl. Microbiol. Biotechnol., 61, 83, 2003. 96. Geisen, R. Multiplex polymerase chain reaction for the detection of potential aflatoxin and sterigmatocystin producing fungi. Syst. Appl. Microbiol., 19, 388, 1996. 97. Farber, P., Geisen, R. and Holzapfel, W. H. Detection of aflatoxigenic fungi in figs by a PCR reaction. Int. J. Food Microbiol., 36, 215, 1997. 98. Geisen, R., Mulfinger, S. and Niessen, L. Detection of Asper­ gillus flavus in wheat by PCR. J. Food Mycol., 1, 211, 1998. 99. Chang, P.-K. et al. Increased expression of Aspergillus parasiticus aflR, encoding a sequence- specific DNA binding protein, relieves nitrate inhibition of aflatoxin biosynthesis. Appl. Environ. Microbiol., 61, 2372, 1995. 100. Yu, J.-H. et al. Conservation of structure and function of the aflatoxin regulatory gene aflR from Aspergillus nidulans and A. flavus. Curr. Genet., 29, 549, 1996. 101. Yu, J.-H. and Keller, N. Regulation of secondary metabolism in filamentous fungi. Annu. Rev. Phytopathol., 43, 437, 2005. 102. Shapira, R. et al. Detection of aflatoxigenic molds in grains by PCR. Appl. Environ. Microbiol., 62, 3270, 1996. 103. Kusumoto, K.-I., et al. Transcript of a homolog of aflR, a regulatory gene for aflatoxin synthesis in Aspergillus parasiticus, was not detected in Aspergillus oryzae strains. FEMS Microbiol. Lett., 169, 303, 1998. 104. Lee, C.-Z., Liou, G.-Y. and Yuan, G.-F. Comparison of the aflR gene sequences of strains in Aspergillus section Flavi. Microbiology, 152, 161, 2006. 105. Cary, J.W. et al. Molecular and functional characterization of a second copy of the aflatoxin regulatory gene, aflR-2, from Aspergillus parasiticus. Biochim. Biophysic. Acta, 1576, 316, 2002. 106. Criseo, G., Bagnara, A. and Bisignano, G. Differentiation of aflatoxin-producing and non producing strains of Aspergillus flavus group. Lett. Appl. Microbiol., 33, 2198, 2001. 107. Chen, R.-S. et al. Polymerase chain reaction-mediated characterization of molds belonging to the Aspergillus flavus group and detection of Aspergillus parasiticus in peanut kernels by a multiplex polymerase chain reaction. J. Food Protect., 65, 840, 2002. 108. Yang, Z.-Y. et al. Detection of aflatoxin-producing molds in korean fermented foods and grains by multiplex PCR. J. Food Protect., 67, 2622, 2004. 109. Manonmani, H.K. et al. Detection of aflatoxigenic fungi in selected food commodities by PCR. Process Biochem., 40, 2859, 2005. 110. Mayer, Z., Farber, P. and Geisen, R. Monitoring the production of aflatoxin B1 in wheat by measuring the concentration of nor-1 mRNA. Appl. Environ. Microbiol., 69, 1154, 2003. 111. Mayer, Z., et al. Quantification of the copy number of nor–1, a gene of the aflaxtoxin biosynthetic pathway by real-time PCR, and its correlation to the cfu of Aspergillus flavus in foods. Int. J. Food Microbiol., 82, 143, 2003. 112. Sweeney, M.J., Pamies, P. and Dobson, A.D.W. The use of reverse transcription polymerase chain reaction (RT-PCR) for monitoring aflatoxin production in Aspergillus parasiticus 439. Int. J. Food Microbiol., 56, 97, 2000.

548 113. Scherm, B. et al. Detection of transcripts of the aflatoxin genes aflD, aflO, and aflP by reverse transcription-polymerase chain reaction allows differentiation of aflatoxin-producing and non-producing isolates of Aspergillus flavus and Aspergillus parasiticus. Int. J. Food Microbiol., 98, 201, 2005. 114. Degola, F. et al. A multiplex RT-PCR approach to detect aflatoxigenic strains of Aspergillus flavus. J. Appl. Microbiol., 103, 409, 2007. 115. Haugland, R.A. and Vesper, S.J. Method for identifying and quantifying specific fungi and bacteria. US Patent and Trademark Office, Patent#6,387,652, 2002.

Molecular Detection of Foodborne Pathogens 116. Doyle, J.J. and Doyle, J.L. A rapid DNA isolation procedure for small quantities of fresh leaf tissue. Phytoch. Bull., 19, 11, 1987. 117. Sartori, D. et al. PCR method for the detection of potential ochratoxin-producing Aspergillus species in coffee beans. Res. Microbiol., 157, 350, 2006. 118. Glass, N.L. and Donaldson, G.C. Development of primer sets designed for use with the PCR to amplify conserved genes from filamentous ascomycetes. Appl. Environ. Microbiol., 61, 1323, 1995. 119. Schimdt-Heydt, M. and Geisen, R. A microarray for monitoring the production of mycotoxins in food. Int. J. Food Microbiol., 117, 131, 2007.

39 Candida

P. Lewis White, Samantha J. Hibbitts, Michael D. Perry, and Rosemary A. Barnes University Hospital of Wales

Contents 39.1 Introduction.................................................................................................................................................................... 549 39.1.1 The Organism................................................................................................................................................... 550 39.1.2 The Incidence, Risk Factors, and Epidemiology.............................................................................................. 550 39.1.3 Pathogenesis...................................................................................................................................................... 552 39.1.4 Spectrum of Disease......................................................................................................................................... 553 39.1.5 Diagnosis Within the Host................................................................................................................................ 553 39.1.6 Detection in Food............................................................................................................................................. 554 39.2 Methods.......................................................................................................................................................................... 558 39.2.1 Sample Preparation and Extraction.................................................................................................................. 558 39.2.2 Detection Procedures........................................................................................................................................ 559 39.2.2.1 Standard PCR.................................................................................................................................. 559 39.2.2.2 Real-time PCR................................................................................................................................. 559 39.3 Conclusions..................................................................................................................................................................... 561 References.................................................................................................................................................................................. 561

39.1  Introduction The genus Candida comprises approximately 200 species and although only a few species have been associated with human disease, an increasing number of susceptible individuals and the emergence of new pathogenic species such as Candida dubliniensis1 make candidal infection a significant problem. Consisting of unicellular eukaryotic cells the organism survives by obtaining nutrients from both living and dead organic matter and Candida often spoils saccharide rich liquids such as fruit juices. The organism is an opportunistic pathogen and a large percentage of infections originate from the commensal reservoir within the patient.2 Candida species are commonly cultured from mucosal membranes and Candida albicans is found in the gastrointestinal tract as part of the normal microbial population. The pathogenic Candida species are unlikely to be found and transmitted in food.3 However, the presence of Candida within foodstuffs may be problematic in debilitated patients particularly those with gastrointestinal mucosal disruption that may allow translocation of the yeast from the GI tract to the blood.4 In a French study it was determined that 11.4% of candidemias originated from within the digestive tract.5 Nevertheless, the focus of Candida within foodstuffs is more of food spoilage than pathogenicity. Accurate and early diagnosis permits the prompt initiation of therapy a factor that may reduce mortality rates and decrease length of stay within hospital.6 It has been estimated that patients with invasive candidal disease spend 10–30 more days in hospital than control patients and this accounts for almost 85% of increased hospital costs.7 Conversely, sensitive

assays with high negative predictive values can exclude candidal infections removing the need for empirical therapy reducing cost and unnecessary exposure to antifungals and subsequent side effects. With an expanding and diverse “at-risk” population Candida species remain the single most prevalent cause of opportunistic human mycoses and invasive candidal diseases confer significant mortality (40–75%) and no less risk of death today than in the previous two decades.8 Candida species are a leading cause of nosocomial bloodborne infection accounting for up to 10% of bloodborne infections acquired in US hospitals where invasive candidal infections are the fourth most common blood stream infection.9 In England, Candida accounts for over 75% of invasive mycoses reported.10,11 Affecting both immuno-compromised and immunocompetent hosts the spectrum of disease ranges from severe irritation due to superficial infections to life threatening disseminated disease requiring intensive medical care. The appearance of superficial infections such as oral candidosis (thrush) may also represent the progression of other underlying conditions such as HIV. In the food industry high levels of fungi within food diminish their appeal by subsequent product degradation leading to economic loss to all parties involved. The accurate diagnosis of both clinical infections and food-spoilage can be difficult and conventional microbiology techniques can be slow and lack both sensitivity and specificity. The development and application of molecular techniques, particularly real-time methodology may alleviate some of these problems. The remainder of this section will 549

550

expand on the review of the biology, epidemiology, pathogenesis, and diagnosis of Candida. However, the subject of Candida and candidosis is extensive and various text books have been written on the topic and should be used for further insight into the field.

39.1.1 The Organism Generally Candida species can be described as dimorphic. The morphology of Candida depends on environmental conditions and the yeast can produce true budding round/oval yeast cells (blastoconidia), pseudohyphae or true hyphal structures, although one species C. glabrata (previously in a separate genus, Torulopsis) rarely produces pseudohyphae or hyphal structures. The cell wall is comprised mainly of carbohydrate, with lesser amounts of protein and lipid forming a multi-layered barrier of mannoprotein (30–40%), β-D-glucan (50–60%) and chitin (0.6–3%).12 β-D-glucan/chitin are essential structural polysaccharides and construct the rigid framework to which mannoproteins are linked providing the cell surface characteristics and properties such as adherence and host immunomodulation.12,13 Colonial appearance is characteristically creamy/white and colonies are generally smooth and wet looking. However, C. albicans possesses the ability to reversibly switch colony surface appearance from smooth to rough and also from creamy/white to opaque a change that corresponds to an alteration in cell shape from round (oval) to elongate. The opaque elongate cells are the ­mating-competent form of C. albicans.14 C. albicans cells can switch without environmental stimulation but pH, oxygen concentration, low intensity treatment with UV and a limitation of zinc in growth medium affect the frequency of “phenotype switching” which has been proposed to enhance pathogenesis of the organism.14–16 The phenomenon of “phenotypic switching” has been used as a laboratory technique to differentiate C. albicans from non-albicans species. When grown on heated blood agar in an atmosphere of 5% CO2 C. albicans colonies grow with an irregular margin that distinguishes them from other Candida species.15 Candidal reproduction is almost entirely asexual by budding which may be multilateral and the formation of chains of connected buds is not uncommon. Candida species prefer aerobic growth conditions, although they will grow in the presence of elevated CO2 but strict anaerobic conditions may inhibit growth.17 Growth is permitted between the temperatures of 20–37°C and within the pH range 2.5–7.5. It appears that 37°C is the optimal growth temperature for more pathogenic species (e.g., C. albicans), albeit all pathogenic strains are capable of growth at this temperature.17 This may be an important factor when considering food spoilage organisms as foodborne pathogens. Growth below 33°C leads to individual yeast cell formation via polar budding, whereas growth at higher temperatures and at neutral pH usually confers germ-tube formation leading to hyphal structures.17 Indeed, the formation of germ-tubes within 3 h when incubated in serum at 37°C is a commonly used laboratory test for determining the identity of C. albicans, although C. dubliniensis

Molecular Detection of Foodborne Pathogens

also possesses this ability.18 The production of large thick walled chlamydospores when grown under a cover slip on corn-meal agar can also be used to distinguish C. albicans. Chlamydospores are formed when grown under nutrient limitation although their function is not known it is proposed they have a role in survival during extreme conditions but are not thought to have a role in pathogenicity.14 C. albicans possesses a diploid genome spanning 1.6 × 107 base pairs and comprising eight pairs of chromosomes. The genome contains genes for 9000 large proteins (>100 amino acids) shares only 60% homology with Saccharomyces cerevisiae and contains more species specific genes which could explain its heightened pathogenticity compared to S. cerevisiae.19–21 Five hundred and sixty seven genes have been deemed essential with an estimated 1400 genes necessary for growth.20 The essential rDNA genes have shown varying copy numbers between both strains and species with some strains of C. albicans possessing over 200 copies making them ideal targets for molecular detection.22

39.1.2 The Incidence, Risk Factors, and Epidemiology Despite improvements in antifungal treatment for invasive candidal infections high mortality rates persist and incidence rates are, at best, described as steady. It is estimated that there will be between 7000 and 28000 hospital acquired candidemias annually in the US of which between 2800 and 11200 patients will die from the disease.8 In 2003 the incidence rate for invasive candidal infections (including candidemias, hepatosplenic candidosis and other deep tissue infections) was 29/100000 US population giving a total 63000 infections per annum.8 Incidence rates for invasive candidal infections vary geographically and tend to be lower in Europe than in the US. In the UK a combined incidence of 2.3–3.1/100000 population has been recorded for England, Wales, and Northern Ireland, although this figure is slightly greater in Scotland (4.8/100000).23,24 In Finland and Norway the incidence is around 2/100000 although the mean annual incidence of candidemia in Iceland between 1991 and 2006 was 4.8/100000 population and was even greater in Denmark (11/100000 population).25–27 Differences in medical practices make it difficult to compare between countries and extrapolating data between countries is not always possible as invasive candidosis is usually a disease of debilitated patients receiving intensive interventional care which varies between countries.4,8 For instance outpatient-diagnosed candidemia is almost three-fold greater in the US than in Spain and reflects the increasing non­inpatient management of chronic diseases in the US.8,28,29 The incidence of invasive candidal infections will also vary between patient groups and in a large European survey candidemia rates were greatest in ICU patients and/or those having received major surgery.25 Rates of candidal infection are greatest in the very young or old, especially neonates and tend to be greater in males than females.8,10 Patient ethnicity also affects the incidence and in two US studies rates have more than quadrupled in black neonates

551

Candida

compared to white neonates and may reflect the increased incidence of premature and/or low weight births within the black population.8,29,30 There are many substantiated risk factors for developing invasive candidal infection, although many are nonspecific and commonly encountered in the critically ill. Some risk factors will vary between patient groups, e.g., neutropenic or nonneutropenic, and some groups have tried to establish predictors of candidemia within specified patient groups, e.g., cardiothoracic ICU.31 However, a more flexible approach may be required when assessing the risk as many risk factors are independently important but also impact on other factors within an individual patient.4 A long-term stay within ICU is a major risk factor as it provides an enhanced possibility of Candida transmission between high-risk patients and the initiation of many practices that increase risk.8 The underlying disease (e.g., malignancy, renal failure, peritonitis, or HIV) may cause immuno-supression and necessary therapy (chemotherapy, corticosteroids, hemodialysis) may accentuate the issue by disrupting the mucosal membranes, providing a route of infection and worsening the immune status. The introduction of medical devices (central venous catheters or mechanical ventilation) may be essential and common practice in ICUs or after surgery but not only provides an entry portal for the organism but also a surface for biofilms to form that may be particularly resistant to treatment.4 The (un) necessary administration of multiple or prolonged antibacterial agents can disrupt the balance of the commensal microbiota promoting fungal colonization that may enter through IV administration of drugs, contaminated foodstuffs or even via contaminated infusate used in total parental nutrition

(TPN) and TPN itself is an independent risk factor. Surgery, particularly involving the gastrointestinal tract is also relevant but other major procedures such as those involving solid organ transplantation place the patient at risk as in most cases ­follow-up immuno-suppressive conditioning will be necessary to prevent rejection of the organ. Candida colonisation is an important independent risk factor. Candida colonization of multiple noncontiguous sites has been linked with increased mortality rates and is often a factor used for initiating pre-emptive therapy.32 Interestingly risk factors for individual Candida species are being proposed. Since the early 1990s the number of Candida glabrata bloodstream isolates has been increasing in the US. This is of particular concern as approximately 20% of isolates will be resistant to fluconazole and almost 10% will be resistant to voriconazole.33 The use of fluconazole has been linked to the increased isolation of C. glabrata but current studies have not established this association and have found age (>60), antibacterial usage, central venous catheters, TPN and ICU residence to be more important.8,34,35 The epidemiology of candidal infections shows geographical variation, although C. albicans remains the most common isolate (Table 39.1). Data from the Artemis Disk Surveillance Programme that collated results from 127 institutions covering 39 countries showed that C. albicans was the most prevalent organism although this figure had decreased by 11% between 1997 (73.3%) and 2003 (62.3%).33 Despite an increasing prevalence in the US (20–24%), C. glabrata levels remained steady between 1997 and 2003, accounting for approximately 10% of isolates globally.8,33 The major increases were seen in the prevalence of

Table 39.1 Geographical Distribution of Candida Species Causing Bloodstream Infections Locations Species (%)

Global

Europe

England and Wales

France

Brisbane, Australia

Brazil

Saudi Arabia

North America

C. albicans

62.3

56

60.1

52.5

53

40.9

53

51

C. glabrata

12.0

14

9.4

11.4

14.5

4.9

7

22

C. tropicalis

7.5

7

3.8

9.5

7.7

20.9

19

7

C parapsilosis

7.3

13

10.6

15.8

10.3

20.5

16

14

C krusei

2.7

2

1.5

4.4

11.1

1.1

2

2

C. guillermondii

0.8

1

0.6

–

2.4

1

<1

C. lusitaniae

0.6

1

0.3

–

–

–

1

<1

–

– –

0.02 –

– –

– –

6.2 –

C. pelliculosa C. dubliniensis C. pseudotropicalis

–

Other

–

Candida spp.

–

Year Reference

2003 1997–9 Pfaller and Tortorano Diekema8 et al.94

0.68

13 1990/9 Lamagni et al.10

–

1.3 5.0

– –

–

3.4

2.6

–

1995 1992–9 2003/4 1996–2004 Richet et al.5 Hope et al.95 Colombo At-Tawfiq97 et al.96

Scotland

52.0 22.7 2.0 11.7 1.0 3.3 2.0

– –

–

–

–

3.0

>2

2.3

–

–

2001/4 Pfaller et al.98

2005/6 Odds et al. 24

552

C. tropicalis (4.6–7.5%) and C. parapsilosis (4.2–7.3%) and there was a slight rise in the number of C. krusei isolates (1.7–2.7%).8 Recent US data has indicated that C. glabrata and C. krusei have become the major etiological agents responsible for candidemia in patients with hematological malignancy whereas C. albicans and C. parapsilosis remain statistically more significant in solid tumour patients.36 An increasing incidence in C. glabrata and C. krusei bloodstream infections is cause for concern as these infections are associated with higher crude mortality rates (45.0 and 55.3%, respectively) when compared to the overall mortality rate (37.9%) for invasive candidal infections. This possibly relates to the increased resistance of these species to fluconazole leading to a delay in initiation of appropriate treatment25 or to a higher rate of co-morbidities in these susceptible patients. European data supports the reduction in the incidence of C. albicans isolates in hematological patients.25 Globally there have been significant increases in the prevalence of other candidal species but these comprise less than 10% of the total number of isolates and are from a range of more than ten species. The top five species causing invasive candidal infections globally has remained constant since 1992: C. albicans, C. glabrata, C. tropicalis, C parapsilosis and C. krusei.8,33 However, this will vary with geographical location and with patient population (Table 39.1). In addition to the life threatening conditions, many more patients will suffer from superficial candidal infections although accurate incidence figures are not known. Surveillance is hampered by clinical diagnosis in a variety of nonmedical settings (e.g., dental surgeries) and by the patient self-medicating with over-the-counter remedies. As with the invasive form of the disease C. albicans is the most common cause of superficial infection and most infections are endogenous. Candida species can be isolated from the oral cavity/GI tract of up to 50% of the population and the genital tract 20% of the female population without symptoms; C. albicans accounts for between 60 and 90% of these isolates.37 Candida parapsilosis is commonly isolated from the skin and has the ability to persist in the hospital environment and infection via contact with family members or healthcare workers has occurred in children and neonates where its ability to form biofilms and colonise catheters is a further concern. Most cases of oral candidosis are caused by C. albicans, with occasional cases caused by C. tropicalis, C. parapsilosis, C. glabrata, and C. krusei.37 Risk factors will vary with type of superficial disease, but underlying immunosuppressive conditions, antibiotic or corticosteroid use and diabetes mellitus may predispose an individual to superficial candidosis. Smoking, dental neglect/ trauma, or poor denture cleanliness/fitting may lead to various types of oral candidosis. Pregnancy, menstruation, certain oral contraceptives, estrogen replacement therapy, and tight clothing have been linked with vaginal candidosis and warm moist areas exposed to maceration can lead to cutaneous candidosis.37

Molecular Detection of Foodborne Pathogens

39.1.3  Pathogenesis The virulence of Candida varies between species and in rodent studies C. albicans is the most potent, followed by C. tropicalis with other species relying on host immunosupression prior to infection.38 As such most studies on pathogenicity have focused on the primary pathogen and C. albicans has several well studied virulence factors. The Candida cell wall is essential for initiating and maintaining the pathogen/host connection and the adherence of Candida to the host is thought to be important in its pathogenicity. Within C. albicans there are eight genes in the ALS family that code for cell well glycoproteins (adhesins) that are adhesive and/or promote flocculation and filamentation. Certain proteins within the family express differing adherence preferences allowing Candida to adhere to different cell types, extracellular matrices or even plastic.38,39 For adhesins to function optimally they need to be situated at the cell surface as is the case for any hydrolytic enzymes or molecules used to avoid host defences.38 The secreted aspartyl proteinase (Sap) is well established as a C. albicans virulence factor and is rarely found outside of the two most pathogenic species C. albicans and C. tropicalis. More recently the use of oxidative stress responses to resist phagocytosis and killing by the host have been shown to be important.38 Within host tissue Candida will take on a mix of morphologies ranging from yeast cells, pseudohyphae through to true hyphal structures and this implies that the traits required for candidal success are present in all morphological forms.40 The formation of germ-tubes in human serum at body temperature indicates that in principle this may be the preferred morphology for Candida within human tissue although in reality varying forms are found demonstrating the effect of environmental pressures.40 Our knowledge of growth regulation of pseudohyphae and true hyphal formation in vivo is limited and it is likely that candidal structure is determined by host susceptibility.40 During infection hyphal structures are more invasive than yeasts and can penetrate deeper into the endothelial tissue expanding the area of tissue damage and allowing increased scope for yeast proliferation.40 However, it cannot be definitively proven that one form of candidal cell is more virulent than the other.40 A range of morphological structures allows C. albicans to adapt to and invade various differing environments and this may be further enhanced by “phenotypic switching” (see above). This reversible process can transform cell morphology, metabolism and surface properties permitting greater scope for flexibility. The white to opaque switch that coincides with a change in cells shape from round/ oval to elongate adapts C. albicans for sexual reproduction that would allow gene transfer and inter-strain propagation of beneficial mutations (e.g., azole resistance). The formation of biofilms on solid phases (including dentures, catheters, artificial heart valves) within the host may also increase resistance to antifungal therapy and this may be linked to an increased production of efflux pumps within the biofilm.41

Candida

Candida contain structures on their cell surface that trigger innate host responses via the toll-like receptors and other recognition molecules.42 Certain mannoproteins on the cell walls can trigger a proinflammatory T-helper (Th1) cytokine response such that acute candidemia may present as profound sepsis. Conversely, exposure to other mannoproteins, dectin and glucans may result in a down-regulatory response associated with anti-inflammatory (Th2) cytokines and persistent fungal infection. This form is more likely to be associated with neutropenia and other causes of monocyte deactivation following immunosuppresssion, trauma/surgery, immaturity, head-injury, or sepsis. The application of this Th1/Th2 paradigm to human infection is far from clear and recent recognition of the role of regulatory T cells (T-reg) and Th17 responses further complicates the picture. Th17 responses are an important cause of immunopathology promoting inflammation, defective pathogen clearance, failure of protective responses to Candida and increased neutrophil inflammatory activity causing tissue damage.43 These are countered by the T-reg response that promotes immune tolerance. Recent evidence suggests Candida can modulate and evade host immunity and use immunomodulatory molecules that enable them to promote protective tolerance leading to colonization and persistence.44

39.1.4 Spectrum of Disease Candida can cause a wide variety of infections and these are summarized in Table 39.2. The most common are superficial infections of the mucosal membranes of the GI and genital tract but the most concerning are the invasive forms that generally occur in predisposed patients. Invasive infections can be localized to an individual organ/region or become systemic (disseminated) where two or more organs become involved or Candida is detected in the bloodstream without organ involvement (candidemia). The organs involved will be dependent on the original site of infection, underlying disorder and the immune status of the patient but all major organs have been infected.

39.1.5  Diagnosis Within the Host Diagnosis of candidal infections is a combination of culture, microscopic, and clinical evidence. In superficial infections culturing Candida is not diagnostic, as the organism is commensal and microscopic evidence of the various candidal forms within swabs/scrapings of lesions or discharge is more significant.37 Positive cultures form superficial sites are not diagnostic for invasive disease but culture from multiple noncontiguous anatomical sites is a predisposing factor. Candida are cultured from the urine of 1–9% of hospital inpatients and is important in immunocompromised patients.45 The diagnosis of invasive candidal infections presents further challenges. Clinical signs may be nonspecific and a combination of tests is required to obtain the correct verdict (see Table 39.2). In 2002 consensus criteria were published to define certainty in diagnosis of invasive fungal infections

553

and these were refined in 2008.46,47 Proven invasive candidal infection was defined as the microscopic or culture detection of any Candida species (pseudohyphae or hyphae) in a needle aspirate or biopsy from a sterile site or recovery of Candida species from the blood.47 However, obtaining a specimen from an internal organ may be too invasive for a critically ill patient. Any deep specimen taken for microscopic testing should also be cultured but occasionally specimens are frozen or embedded in paraffin prior to culture rendering the organism nonviable. Multiple blood cultures should be taken but 50% of patients with autopsy proven invasive candidal infection are blood culture negative.45 Lytic blood culture systems using large volumes of blood (20 ml) with extended incubation periods will improve the sensitivity but this is not routinely performed in most centres. Depending on blood culture system most detectable candidemias will be grown within 3 days; however extended incubation may be required. Combining this with the time required for identification (germ-tube or chlamydiaspore production, API ID32C, Candida AUXACOLOR or Chromagar) introduces a 72–168 h delay before accurate species diagnosis can be made and this may have consequences for treatment, particularly for azole resistant strains. The limitations of classical tests restricts the number of diagnosed proven invasive candidal infections and clinical signs such as the CT detected bull’s eye lesion in liver or spleen or retinal exudates are often combined with predisposing factors to initiate pre-emptive or empirical therapy. One such approach is the colonization index, where the ratio of the number of Candida culture positive sites to the number of sites cultured is used to start pre-emptive therapy. This technique has shown a significant reduction in the incidence of invasive candidal infections.48 An alternative risk strategy is to calculate a “Candida score,” where the presence of independent risk factors (surgery, multifocal colonisation, TPN, and severe sepsis) for ICU patients are collated to generate a threshold above which early therapy is commenced.32 Two major drawbacks of this approach are that many of the risk factors are common, particularly in ICU patients and many of the clinical signs are non-specific. This leads to the unnecessary use of antifungal therapy, exposure to side-effects, potential to promote resistance and additional expense. The introduction of nonculture diagnostic techniques has the potential to drastically enhance the detection of both invasive and superficial candidal disease. Serological methods targeting both candidal antibodies and cell wall antigens have been widely applied. The main benefit of both antigen and antibody detection is quality control that is achieved by the availability of standardized commercially available kits. Both latex agglutination and ELISA kits are available although the former has a reduced sensitivity.49 For a more expansive review of fungal cell wall immunodiagnostics see the recent article by Kedzierska et al.50 Table 39.3 collates some of the published articles directly comparing PCR and immunodiagnosis.

554

Molecular Detection of Foodborne Pathogens

Table 39.2 The Range of Candidal Disease Type Superficial

Clinical Manifestation

Clinical Form

Symptoms/Signs

Oral candidosis

Pseudomembranous candidosis (oral thrush) Acute atrophic candidosis Chronic atrophic candidosis Angular cheilitis Chronic hyperplastic candidosis

Genital candidosis

Vaginal candidosis

White raised lesions/plaques—removal leaves sore erythematous (bleeding) surface Pain, swelling, tongue depapillation, restricted tongue movement Asymptomatic, erythema, oedema, angular cheilitis Soreness, erythema, and fissuring at the corners of the mouth. Asymptomatic, non removable lesions/plaques on buccal mucosa, can lead to malignancy. Vaginal/vulval itching/burning with possible discharge. Erythema with fissuring, plaques, dysuria. Soreness, itching, lesions and erythema, occasional fissuring Lesions within skin folds leading to vesicles that may rupture to form erythematous regions. Swelling, erythema and pain of nail fold, marks appear on the nail before pitting and detachment of the nail. Widespread cutaneous and/or mucosal lesions.

Penile candidosis Cutaneous candidosis Candida paronychia

Invasive

Chronic mucotaneous candidosis GI candidosis

Pulmonary candidosis Central nervous system candidosis Candida heart infections

Oesophagitis Gastric candidosis Intestinal candidosis Candida meningitis Brain abscess/ encephalitis

Renal candidosis Candida peritonitis Candida endopthalmitis Systemic

Candidemia Acute disseminated candidosis Chronic disseminated candidosis

Oesophadynia, dysphagia, ulceration Often asymptomatic, ulceration common Ulceration. Nonspecific signs. Bilateral nodular infiltrates. Identical to bacterial meningitis CT/MRI detected lesions Nonspecific signs: Chest pain, pericardial effusion or rub, Fever, weight loss, fatigue, enlarged spleen, murmurs. Large vegetations, embolisation and endopthalmitis, Nonspecific signs: Abscess formation, Lumbar/abdominal pain, fevers, rigors. Nonspecific signs: abdominal pain, fever, nausea, vomiting. Nonspecific signs: Blurred vision, pain, floaters. Typical signs: yellowwhite retinal lesions. Nonspecifc signs: Fever, rigors, hypertension Nonresponsive fever, cutaneous lesions, endopthalmitis. Leading to many of the conditions described above. Nonresponsive fever, abdominal pain, enlarged liver and/or spleen, lesions on liver or spleen, bullseye lesion, raised alkaline phosphatase levels

Source: Summarized from Richardson, M.D. and Warnock, D.W., Fungal Infections: Diagnosis and Management 3rd edn, Blackwell Science Ltd., Oxford, London, 2003.

There have been a large number of publications describing the successful design and application of PCR as an adjunct test in the diagnosis of Candida and a selection are summarized in Table 39.4. Generally sensitivity and specificity is high although no consensus has been reached regarding the optimal specimen, extraction technique or amplification system and interpretation of PCR results without clinical or microbiological support is difficult. The application of molecular diagnosis of candidal infections within the host will not be covered in detail in this chapter. Instead some of the potential benefits of the many clinically applied PCR methods will be discussed in reference to detecting Candida as a whole.

39.1.6  Detection in Food Much of the conventional detection of Candida in food involves the isolation and enumeration of yeasts via culture using low pH media such as potato dextrose or malt extract agar.3 In practice this is a tedious process involving sample preparation and dilution, several cultural enrichment stages, enumeration, isolation, and identification by morphological, immunological, or biochemical characteristics.3,51 These procedures can take considerable time and although considered cost effective the labor involved will add significant expense. The food matrix will affect recovery efficiency and for solid media extensive violent disruption

555

Candida

Table 39.3 Comparison of Clinical Nonculture Diagnostic Techniques Positivity Rate(%)

Mouse model

β-D-Glucan

PCR

Population Day 7.5

7.1

Day 8 Day 9

60 100

ELISA

DFI

LA 100 – –

Day 7.5

100

–

Day 7.5

– –

– –

– – 47d

– –

N = 27

88

47

N = 14

95

–

75

N = 12

100

–

N = 24

75

54

a b c d e f g h

41c

75e

Reference Uno et al.99, a – –

–

Alam et al.100, b

–

25

White et al.80, f

–

–

58

Ahmad et al.75, g

–

–

21

Sakai et al.70, h

Serological tests: Mannan, Unimedi-Candida. β-D-Glucan, β-Glucan Wako. Serological tests: Mannan, Platelia Candida Ag. Candida antibody, Platelia Candida Ab/Ac/Ak kit. β-D-Glucan, Fungitell. Mannan performance alone. Anti-Candida antibodies performance alone. Combined performance. Serological tests: Mannan, Platelia Candida Ag and Pastorex Candida. Serological tests: Candida antibody, Cand-Tec test. Serological tests: β-D-Glucan,Fungitec G test MK. Candida antibody, Cand-Tec test.

is required to dislodge the yeast that will be adhered to the food. Membrane filtration may be required to enhance detection of low cell numbers and can be applied to liquids. Dilution and plating must be performed immediately to avoid cell sedimentation leading to a loss in sensitivity.3 Furthermore, the detection of pathogenic or spoilage strains may be masked by the large number of background organisms.52 Similar to clinical mycology most yeast will grow within 3 days although an extended incubation of 5 days enhances detection. Unlike clinical mycology plates will be incubated between 25 and 28°C and this is important when considering a foodborne organism as a possible pathogen that must be able to tolerate growth at 37°C. Once cultured identification of the Candida species can then be attempted by studying morphological differences both colonial and microscopic or by biochemical tests utilizing differences in metabolism. Candida species will vary in their ability to ferment a range of sugars and their ability to assimilate different nitrogen and carbon sources and these differences represent the basis of both commercial and inhouse identification systems.53 The development of PCR technology has revolutionized many clinical fields (e.g., virology) and its application in scientific disciplines is widespread. Unlike phenotypic profiles molecular detection will remain static in the short term and is uninfluenced by environmental conditions.54 With yeast playing important roles in many productive processes (e.g., wine fermentation) it is important to maintain population levels and prevent the introduction of spoilage organisms.55 The use of PCR in the food industry has improved turnaround times and accuracy of results. The

use of molecular typing methods can be used to trace the source of organisms and their route of entry into the food chain permitting the application of adequate preventative measures. A process that would be almost impossible to achieve by conventional means.54 The additional conventional limitation of the viable but nonculturable microbial population can also be targeted by applying molecular technology.54 One major weakness with molecular detection within food remains its reliance on cultural enrichment to achieve adequate performance. Currently this is a necessity as nucleic acid extraction systems struggle to remove much of the associated food debris or inhibitory compounds within.51,56 Soft cheeses and calcium ion in milk have shown inhibitory properties as have ferric ammonium citrate, bile salts, esculin and acriflavin, and several ionic detergents.51 Conversely, the enrichment process generating larger quantities of biomass can lead to the extraction of inhibitory DNA concentrations, 56 although the amount of culture can be quantified prior to commencing the process. As PCR amplifies DNA it is also unable to distinguish between viable and nonviable organisms and may not be able to demonstrate the efficacy of food-treatment processes leading to false positive results. Messenger RNA is an indicator of viability and its detection by reverse transcriptase (RT) PCR or nucleic acid sequence based amplification (NASBA) is possible, although the stability of RNA through the ubiquitous nature of RNAses is a concern.56 Table 39.4 summarizes a range of PCR methodology used for detecting Candida or fungi in a clinical setting. Although these assays have been developed principally for clinical

556

Molecular Detection of Foodborne Pathogens

Table 39.4 A Selection of Published PCR Protocols Suitable for Detecting Candida Target Region

PCR Type

Detection Limit

Detection Range

Reference

ITS region 18s rRNA

Semi-nested Conventional

One organism/ml 4.5 CFU/ml

Calb, Ctrop, Cpara, Cglab Pan Fungal

Ahmad et al.75 Sakai et al.70

28s rRNA

Real-time

<10 organism/reaction

Bašková et al.84

28s rRNA 18s rRNA

Real-time Real-time

15 copies/reaction 10 CFU/ml

Asp, Calb, Cglab, Ckrus, Cguill, Ctrop, Cpara, Ckef, Clus, Cdub panfungal panfungal

Vollmer et al.83 Jordanides et al.89

ITS region

Semi-nested

1 Fg DNA

panfungal

Bagyalakshmi et al.76

18s rRNA

Southern blot

4 CFU/ml

panfungal

Van Burik et al.79

ITS region ITS region ITS region

PCR-ELISA PCR-MLH PCR-sequence

2 CFU/ml 2–20 CFU NA

panfungal panfungal panfungal

Badiee et al.101 Hendolin et al.102 Lau et al.103

End-point fluorescence Real-time

One cell per 2 µl sample

Calb, Ctrop, Cpara, Cglab, Ckrus

Shin et al.104

2 CFU/ml

Calb, Ctrop, Cglab, Cpara, Ckrus

Klingspor and Jalal90

ITS 2 region 18s rRNA

Real-time Southern Blot

5–10 CFU/ml 1 CFU/ml

Calb, Cglab, Cpara, Cdub, Ckrus, Ctrop Calb, Ctrop, Cpara, Cglab, Ckrus

Schabereiter-Gurtner et al.81 Einsele et al.77

18s rRNA

PCR-ELISA

5 CFU/ml

Calb

Löffler et al.105

ITS 2 region

Real-time

5 CFU/ml

Calb, panfungal

Maaroufi et al.86

P450 LDMG

RFLP

200 Fg DNA

Morace et al.72

NASBA

1–10 CFU/ml

Widjojoatmodjo et al.92

Real-time

NA

Calb, Cpara, Ctrop, Cguill, Ckrus, Cglab, Ckef Candida species, Calb, Ckrus, Ctrop, Cglab, Clus Calb, Ckef, Cpara, Cglab, Ctrop, Ckrus

Dot blot

10–50 CFU/ml

Calb

Arancia et al.78

Auto capillary Electrophoresis Real-time

NA

Yeasts

Chen et al.106

1–5 CFU/ml

Calb, Cdub, Cglab, Ctrop, Ckef, Ckrus, Cpara

White et al.80

ITS 2 region 18s rRNA

18s rRNA ITS 2 region Cahsp70 ITS 2 region 18s rRNA

Guiver et al.87

18s rRNA and ITS region ITS region

Real-time

0.22–2.3 copies/ml

Calb, Cpara, Cglab, Ctrop, Cdub, Ckrus

McMullan et al.82

Multiplex PCR

2 CFU/ml

Carvalho et al.107

RNase P RNA gene

Multiplex real-time PCR

50 copies

Calb, Cglab, Cpara, Ctrop, Ckrus, Cguill, Clus, Cdub Calb, Cdub, Cfam, Cglab, Cguill, Ckrus, Cpara, Ctrop

Innings et al.85

Abbreviations: Asp: Aspergillus sp.; CaHSP70: C. albicans heat shock protein 70; Calb: C. albicans; Cdub: C. dubliniensis; Cglab: C. glabrata; Cguill: C. guillermondii; Cfam: C. famata; Ckef: C. kefyr; Ckrus: C. krusei; Clus: C. lusitaniae; Cpara: C. parapsilosis; Ctrop: C. tropicalis; MLH: mutliplex liquid hybridization; NASBA: nucleic acid sequence based amplification.

diagnosis the amplification procedure can also be applied to screening foodstuffs. However, specimen types, enrichment processes and DNA extraction techniques will need to be optimized for the particular food and for removal of any

interfering compounds within. This does raise the issue of differences in etiology between the food industry and clinical diagnosis. Clinical assays designed to detect specific pathogenic species may not be sufficiently diverse to be applied

557

Candida

Figure 39.1  Sequence alignment of the 18S rRNA gene (partial sequence) from C. albicans, A. fumigatus and human.

to screening foodstuffs possibly containing environmental strains. A panfungal approach may be better suited but this comes with its own limitations. Figure 39.1 shows the sequence similarity of the small rRNA subunit of C. albicans, Aspergillus fumigatus and Homo sapiens; a gene often used for PCR because of its multicopy nature. Its similarity across eukaryotic organisms has implications when designing a panfungal assay, as oligonucleotides will be based on the conserved regions of this gene which if incorrectly

selected may also bind to many animal genes thus affecting the performance of a screening PCR assay. A commercial PCR system for the detection of yeast and mold in food is available. The BAX® system allows a standardized quality controlled approach for the automated detection of fungi in a variety of food, including cheese, with or without enrichment. However, to achieve the optimal detection limit (10–50 cfu/g) a 44-h enrichment is required although results will be returned after 48 h.

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39.2  Methods 39.2.1 Sample Preparation and Extraction For molecular methods to achieve their full potential in detecting foodborne pathogens they must exclude the need for enrichment culture. The obvious solution is a direct approach, where a fraction of food is used for nucleic acid extraction. However, with composition of food varying greatly no universal procedure is ever likely to exist. Furthermore, for large scale production facilities the question as to the amount of test specimen and frequency of screening to achieve optimal detection of pathogen or food spoilage agent will vary on the foodstuff, the environmental conditions, treatment regimes and growth rate of the target organism. Direct extraction techniques from liquids should be easier to implement. Collection filters have been used to retain a variety of Candida species from water samples prior to DNA extraction and direct extractions have been used to detect yeast in orange juice and wine.55,57–62 Fungal RNA has also been extracted from yogurt and other pasturized dairy products, although an initial sodium chloride dilution and centrifugation was applied.63 For solid foods direct extraction is more problematic. Rapid separation and concentration techniques have been applied to bacterial food pathogens. Using the principle of buoyant density gradient centrifugation Fukushima et al. were able to detect 12 bacterial pathogens from 13 food types, including meat, bread, fish, and shellfish with a detection limit of 101–103 cfu/g for real-time PCR.52 The process involved mixing 25 g of sample with tween20-BPW, prior to homogenization in a stomacher. Filtered homogenates were then centrifuged at low speed and the upper layer centrifuged at high speed. The pellet was retained and washed before being put through flotation and sedimentation buoyant density centrifugation to purify the pathogens and resulting in a theoretical 250-fold increase in concentration.52 Other physical mechanisms for separation and concentration include aqueous polymer two phase partitioning, centrifugation, filtration, and ultrasound treatments.51 The first of these exploits the surface properties of the organism and interactions with two immiscible polymer solutions. As the polymers separate the pathogenic cells will be attracted to one of the fractions depending on whether the cell is hydrophilic or hydrophobic. Unfortunately some cells will congregate at the interface along with most food debris. The method is also limited to small sample volumes and often requires dilution of food specimens to allow efficient separation.51 The role of centrifugation and filtration is self-explanatory. Filters separate the organism from the food by size, although filter clogging is a problem. In principle high speed centrifugation pellets any cells enabling the supernatant (containing specimen) to be decanted. In practice food debris will pellet with the cells, although a dramatic reduction in initial specimen volume is achieved. More complex centrifugation methods have been developed involving various treatments to remove food debris.51 Ultrasound treatment works by clustering cells that are

Molecular Detection of Foodborne Pathogens

allowed to separate by sedimentation and it has been successfully applied to the separation of yeast cells in PBS but is restricted to small volumes and studies in solid matrices have not been performed.51,64 Alternatively, adsorption separation and concentration methods could be used. These work by attaching an antibody, bacteriophage or molecule with an affinity for the pathogen to a solid matrix. Specific target antigens on the surface of the pathogen will bind to the matrix that can then be separated from the food specimen by a variety of techniques. Problems arise due to the presence of the food preventing the attachment of the pathogen to the matrix and by inhibiting its release once attached.51 Once the organism has been separated, concentrated and/ or cultured the basic nucleic acid extraction technique is relatively straightforward. Lysis of the cell wall is essential and can be performed enzymatically, chemically or mechanically. Early manuscripts describe the use of zymolase to form spheroplasts but this should be replaced by lyticase and many clinical studies use recombinant lyticase at great expense. The reason is zymolase originates from the yeast Saccharomyces cerevisiae and its use is a potential source of fungal DNA contamination.65,66 Proteinase K is another fungal derived enzyme commonly used in nucleic acid extraction procedures. Originating from the environmental mold Tritirachium album proteinase K usage is a potential source of contamination if panfungal amplification systems are used. The use of zymolase or lyticase has been used in studies comparing different extraction procedures from fungal cultures.67 In this study the authors compared several commercial extraction kits and found the Qiagen Qiamp tissue kit generated a similar sensitivity to their in house extraction procedure detecting 1–10 cfu but took half the time. In a recent study six alternative methods were compared to the Qiagen/ Lyticase approach (gold standard reference) with respect to detection limit, cost, and completion time.68 Interestingly, in terms of overall performance the authors found the YeaStar genomic DNA kit (Zymo research) was optimal. Taking 1 h to complete with only 15 min hands-on time the zymolase/ spin column approach had the same detection threshold as the reference method but Ct values were significantly better indicating improved sensitivity at less than half the cost and completion time.68 No zymolase linked contamination was found in negative extraction controls and is probably a result of their accurate Candida probe specificity, however, the presence of contaminating S. cerevisiae DNA cannot be excluded. An alternative to enzymatic lysis is mechanical disruption by bead-beating. The use of specialized bead-beaters has been described but the purchase of specialized equipment is not necessary. Simply vortexing the candidal pellet with the equivalent of 20–30 µl acid washed glass beads (180 μm) for 30 sec to 1 min is satisfactory to lyse the candidal cell. This can be combined with most DNA purification procedures and has been combined successfully with manual and automated extraction.68,69 Loeffler et al.69 described the combined use of bead-beating and the Roche MagNA Pure LC automated

Candida

extractor to isolate DNA from 23 yeast and mold species and in terms of performance it functioned as well as the gold standard reference method above.69 A variety of candidal DNA extraction methods are currently available and in our experience simple chemical lysis by guanidine thiocyanate or hot NaOH/SDS is sufficient to lyse the cell wall and is sufficient for Candida culture but may suffer a loss in sensitivity if a direct specimen extraction is being performed.

39.2.2  Detection Procedures A wide range of molecular detection procedures have been described and some are presented in Table 39.4. In this section they will be divided into conventional and real-time PCR and the benefits and limitations described. 39.2.2.1 Standard PCR Conventional PCR describes the use of traditional thermal block to amplify the target of interest. On completion (approx. 3 h) the user is unaware if the reaction is successful and has to perform a downstream process to confirm this. In its simplest form this involves the combination of PCR amplification with identification via agarose gel electrophoresis.70 In terms of sensitivity this is satisfactory for culture, but the system is limited regarding specificity as the amplified product is identified solely by its size. This has lead to a variety of analysis techniques many of which have been applied to Candida. Single strand conformation polymorphism (SSCP) can be used to distinguish Candida species by the sequence variation within a PCR product.71 Targeting multicopy genes (e.g., rRNA) increases the possibility of variation within a strain and will further increase species level discrimination. Denatured single strand DNA will take on a particular conformation dependent on its sequence. Different conformations result in different degrees of retention within a gel and the generation of species level profiles. In principle this is simply a more advanced form of gel electrophoresis but the major drawback are difficulties in preparing and running the gel. PCR restriction fragment length polymorphism (PCRRFLP) is the use of restriction enzymes to digest PCR products, the resulting fragments are separated by gel electrophoresis and the pattern produced should permit genus/species/ strain discrimination. It has been applied to P450 lanosterol 14α-demethylase gene and ITS regions for the identification of Candida species.72,73 Another method using restriction enzymes is amplified fragment length polymorphism (AFLP). The enzyme is used to digest genomic DNA after which PCR adaptors are ligated to fragments and specificity is maintained through pre-selective and selective PCR. Its use has been applied for the discrimination of nine candidal species.74 All the methods above are well suited to use where culture has been achieved, it is unlikely that they will be successfully applied to systems where cultivation is excluded as they are reliant on the amplification of an intensely visible PCR product. Where biomass concentration is less this may not be achieved and even the amplification of a weaker amplicon

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may result in no/mis-identification. In such cases nested or semi-nested PCR have been applied.75,76 Incorporating two rounds of PCR drastically increases sensitivity, but also the possibility of contamination. By far the most accurate technique is PCR combined with Southern blotting.77–79 On completion of PCR the product is transferred to a nylon filter, denatured and hybridized with a specific probe that nowadays can be visualized via a chemical reaction. Sensitivity is sufficient that culture is not necessary and specificity is maintained by well designed primers and probe. All the methods above suffer from the same major drawbacks. A delay in result production due to the necessity of excessive post-amplification handling that also increases the hands-on time, required expertise and the opportunity for false positive results due to contamination. With the development of real-time platforms addressing these problems much of the above technology has been superseded. 39.2.2.2 Real-time PCR Real-time PCR has been in place approximately a decade and a wide range of platforms are available. With amplification taking less than 1 h on certain systems compared to 3 h by conventional PCR and result determination during rather than after the process the benefits are obvious. With the removal of post-amplification handling and the incorporation of uracyl-DNA-glycosylase (UDG) to inactivate previously amplified PCR products post-extraction contamination should not be a problem and real-time PCR should provide greater specificity over its conventional counterpart. Its basic form uses the fluorescent dye SYBR green. As amplification occurs SYBR green binds to double stranded DNA, the amount bound increases proportionally with the amount of amplicon and on passing the background fluorescent level a crossing point (Ct value) is generated that is dependent on the starting amount of DNA target. Unfortunately SYBR green will bind to any form of double stranded DNA including nonspecific amplification and oligonucleotide dimers generating a positive signal. Although melt-curve analysis should resolve this, the introduction of Candida specific Cy5 labeled bi-probes have been used to increase sensitivity and specificity.80,81 When bound to the PCR product the probe accepts energy from SYBR Green and emits a specific signal. The use of bi-probes has been largely replaced by Hydrolysis (Taqman) and hybridization (FRET) probes. Hydrolysis probes are dual-labeled oligonucleotides with a reporter dye at the 5′ end and quencher at the 3′ end. The probe binds very tightly to the target sequence prior to PCR extension and as the Taq polymerase extends from the primer it fragments the oligonucleotide separating the reporter and quencher dyes releasing a signal. A large number of Candida specific hydrolysis systems have been described.82–87 An alternative is the use of hybridization probes.88–90 Two fluorescently labeled probes are designed to bind to the target sequence in a head to tail arrangement. During anneal the probes bind and the oligonucleotide labeled with 3′ fluorescein donates excitation energy to the second oligonucleotide

560

labeled with 5′ LC red 640 and a fluorescent signal is emitted.88 The probes are dislodged during PCR extension and it is possible to perform melt-curve analysis allowing sequence variation to be detected. A process that has been utilized to detect point mutations responsible for fluconazole resistance in C. albicans.91 For both hydrolysis and hybridization systems the Ct value is dependent on initial target DNA concentrations and so can be used to quantify PCR reactions (Figure 39.2). Candidal RNA has also been targeted via NASBA and RT-PCR.63,92 Both RT-PCR and NASBA Candida methods are endpoint analysis system suffering similar limitations to conventional PCR. To our knowledge no real-time NASBA methods utilizing beacon technology have been described. However, RT-PCR has been combined with real-time detection but only after the synthesis of cDNA.55,63 It is possible to perform the reverse transcription and real-time PCR in a single tube and this should be performed to avoid the problems mentioned. Below a detailed quantitative real-time PCR system is described. It is based on a qualitative bi-probe assay (SYBR Green with Cy-5 probe) designed to detect, but not differentiate the main pathogenic species of Candida (i.e., C. albicans, C. dubliniensis, C. glabrata, C. kefyr, C. krusei, C. parapsilosis, and C. tropicalis) using the Idaho Technologies LightCycler system.80 Due to sequence variations the original assay was found to have a variable detection limit when testing certain species, particularly reduced for C. glabrata.93 Furthermore, the development of new real-time technologies lead to the modification of the assay for use on more widely used improved platforms (e.g., Roche Light-Cycler or Applied BioSystems TaqMan) and was multiplexed to allow the differentiation of C. glabrata and C. krusei. This assay uses the same pan-fungal primers (L18F, 5´-CTCGTAGTTGAACCTTGG; L18R, 5´-GCCTGCTTTGAACACTCT) that target two conserved regions encompassing a variable region within the 18S rRNA gene, resulting in the production of a 140-bp amplicon that corresponds to the nucleotide positions of 620–760 in the 18S rRNA gene of C. albicans. The amplified product is

Molecular Detection of Foodborne Pathogens

then detected by hydrolysis probes (generic Candida probe, 5′-FAM ATC TTT TTG ATG CGT ACT GGA CCC TG – BHQ1, C. glabrata probe, 5′-JOE GGC TAA CCC CAA GTC CTT GTG GCT T – BHQ1 and C. krusei 5′-ROX TAC CTA TGG TAA GCA CTG TTG CGG C – BHQ2) that bind to a Candida-specific sequence within the PCR amplicon. Protocol. (1) Extract DNA from Candida isolates using recombinant lyticase (Sigma-Aldrich) and the QiaAmp tissue kit (Qiagen).67 Note: For a cheaper alternative simply vortex the harvested Candida culture with 180 μm acid washed glass beads (Sigma-Aldrich) for 30 sec, wash with molecular grade water and transfer washings to the Qiamp tissue kit. If enhanced sensitivity is required when precipitating DNA with ethanol, the solution is incubated on ice for 30 min to increase the yield. Secondly, after the elution of the DNA, YM-100 microconcentrators (Millipore) are used to increase the DNA concentration. All reagents are filter-sterilized through 0.2 µm filters before use. (2) Prepare PCR mixture (20 µl) consisting of 2 µl Light-Cycler FastStart DNA Master hybprobe master-mix (Roche) [containing dNTPs, Faststart Taq DNA polymerase, 10 mM MgCl2], 2.0µl each of L18F and L18R primers (final concentration 750 nM), 2.0 µl Candida probe (final concentration 400 nM) 2.4 µl of 25mM MgCl2, up to 9.6 µl template DNA and molecular grade water as required. Note: The probe can be monoplex or multiplex. (3) Conduct PCR amplification on a Roche Light-Cycler, Corbett Rotorgene or Applied Biosystems TaqMan as follows: one cycle of 95°C for 10 min; 50 cycles of 95°C for 15 sec, 62°C for 30 sec with a single acquisition to the required channels (FAM-generic, JOEglabrata and ROX- krusei), one cycle 40°C for 30 sec. Typical results are presented in Figure 39.2.

Figure 39.2  Amplification of DNA extracted from varying concentrations of Candida krusei using a species specific hydrolysis probe.

561

Candida

Note: The generic assay has been optimized for Roche Light-Cycler and may require further optimization for other platforms. The multiplex assay has only been performed on the Corbett Rotorgene and different fluorophores are required for use on the Roche LightCycler. The Candida assay specifically recognizes C. albicans, C. kefyr, C. krusei, C. dubliniensis, C. tropicalis, C. parapsilosis, and C. glabrata but not C. guillermondii, C. lipolytica, C. famata, A. fumigatus, A. niger, S. cerevisiae or Cryptococcus neoformans. Although all the species above produce an amplicon with primers L18F and L18R only C. albicans, C. kefyr, C. krusei, C. dubliniensis, C. tropicalis, C. parapsilosis, and C. glabrata produce a probe signal. The assay has a sensitivity of 5 C. albicans cfu per ml of spiked whole blood.80





39.3  Conclusions With Candida not widely associated as a foodborne pathogen, the molecular detection of this genus mainly focuses on discovering its presence within the host where infection, usually endogenous can have serious consequences. However, even though it does not cause the typical illnesses associated with food-borne infections its presence in food could be problematic to certain populations and invasive candidal disease has been linked to translocation from the gut. Immunosupressed patients with predisposing factors for developing invasive disease, particularly those with damage to the mucosa of the GI tract need not be exposed to excessive numbers of exogenous Candida within food and drink. Combining this with losses due to food-spoilage and the limitations of conventional monitoring techniques it highlights the need for a rapid but accurate screening system. Interestingly the problems within clinical diagnosis and the food industry appear similar where culture techniques are slow and lack sensitivity. The use of PCR in both fields is governed by the extraction procedure. Clinically efficient direct extraction systems have been developed but cannot be applied to the food industry. It is unlikely that an individual DNA extraction process for all the different food types will exist, although DNA extraction from the organism could be standardized and various upstream processes attached depending on the initial food. PCR technology, provided it is well designed and optimized can be applied to either field although the use of broader detection range assays (pan-candidal/fungal) may be better suited. Until DNA extraction systems can cope with the matrices of difficult food separation, concentration techniques will need to be applied. Indeed, a combination of culture and rapid molecular ID may still be beneficial.

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Molecular Detection of Foodborne Pathogens 42. Netea, M.G. et al. Immune sensing of Candida albicans requires cooperative recognition of mannans and glucans by lectin and toll-like receptors. J. Clin. Invest., 116, 1642, 2006. 43. Zelante, T. et al. IL-23 and the Th17 pathway promote inflammation and impair antifungal immune resistance. Eur. J. Immunol., 37, 2695, 2007. 44. Romani, L. and Puccetti, P. Controlling pathogenic inflammation to fungi. Exp. Rev. Anti-Infect. Therapy, 5, 1007, 2007. 45. Denning, D.W., Kibbler, C.C. and Barnes, R.A. British Society for Medical Mycology proposed standards of care for patients with invasive fungal infections. Lancet Infect. Dis., 3, 230, 2003. 46. Ascioglu, S. et al. Defining opportunistic invasive fungal infections in immunocompromised patients with cancer and hematopoietic stem cell transplants: an international consensus. Clin. Infect. Dis., 34, 7, 2002. 47. De Pauw, B. et al. Revised definitions of invasive fungal disease from the European Organization for Research and Treatment of Cancer/Invasive Fungal Infections Cooperative Group and the National Institute of Allergy and Infectious Diseases Mycoses Study Group (EORTC/MSG) Consensus Group. Clin. Infect. Dis., 46, 1813, 2008. 48. Piarroux, R. et al. Assessment of preemptive treatment to prevent severe candidiasis in critically ill surgical patients. Crit. Care Med., 32, 2443, 2004. 49. White, P.L., Archer, A.E. and Barnes, R.A. Comparison of non-culture based methods for detection of systemic fungal infections, with an emphasis on invasive Candida infections. J. Clin. Microbiol., 43, 2181, 2005. 50. Kedzierska, A. et al. Current status of fungal cell wall components in the immunodiagnositcs of invasive fungal infections in humns: galactomannan, mannan and (1-3)-β-D-glucan. Eur. J. Clin. Microbiol. Infect. Dis., 26, 755, 2007. 51. Benoit, P.W. and Donahue, D.W. Methods for rapid separation and concentration of bacteria in food that by-pass time-consuming cultural enrichment. J. Food Prot., 66, 1935, 2003. 52. Fukushima, H. et al. Rapid separation and concentration of food-borne pathogens in food samples prior to quantification by viable cell counting and real-time PCR. Appl. Environ. Microbiol., 73, 92, 2007. 53. Kurtzman, C.P. et al. Methods to identify yeasts. In Yeasts in Food, Boekhout, T. and Robert, V. (eds.). Behr’s Verlag Publishers, Germany, 2003. 54. Van der Vossen, J.M.B.M. et al. PCR methods for tracing and detection of yeasts in the food chain. In Yeasts in Food, Boekhout, T. and Robert, V. (eds.). Behr’s Verlag Publishers, Germany, 2003. 55. Hierro, N. et al. Real-time Quantitative PCR (QPCR) and reverse transcription QPCR for detection and enumeration of total yeasts in wine. Appl. Environ. Microbiol., 72, 7148, 2006. 56. Scheu, P.M., Berghof, K. and Stahl, U. Detection of pathogenic and spoilage mirco-organisms in food with the polymerase chain reaction. Food Microbiol., 15, 13, 1998. 57. Brinkman, N.E. et al. Evaluation of a rapid, quantitative realtime PCR method for enumeration of pathogenic Candida cells in water. Appl. Environ. Microbiol., 69, 1775, 2003. 58. Ros-Chumillas, M. et al. Evaluation of a rapid DNA extraction method to detect yeast cells by PCR in orange juice. Food Control, 18, 33, 2005. 59. Martorell, P., Querol, A. and Fernandez-Espinar, M.T. Rapid identification and enumeration of Saccharomyces cerevisiae cells in wine by real-time PCR. Appl. Environ. Microbiol., 71, 6823, 2005.

Candida 60. Delaherche, A., Claisse, O. and Lonvaud-Funel, A. Detection and quantification of Brettanomyces bruxellensis and “ropy” Pediococcus damnosus strains in wine by real-time polymerase chain reaction. J. Appl. Microbiol., 97, 910, 2004. 61. Phister, T.G. and Mills, D.A. Real-time PCR assay for detection and enumeration of Dekkera bruxellensis in wine. Appl. Environ. Microbiol., 69, 7430, 2003. 62. Ibeas, J.I. et al. Detection of Dekkera brettanomyces strains in sherry by a nested PCR method. Appl. Environ. Microbiol., 62, 998, 1996. 63 Bleve, G. et al. Development of reverse transcription (RT)PCR and real-time RT-PCR for rapid detection and quantification of viable yeasts and molds contaminating yogurts and pasteurised food products. Appl. Environ. Microbiol., 69, 4116, 2003. 64. Limaye, M.S. and Coakley, W.T. Clarification of small volume microbial suspensions in an ultrasonic standing wave. J. Appl. Microbiol., 84, 1035, 1998. 65. Loeffler, J. et al. Contaminations occurring in fungal PCR assays. J. Clin. Microbiol., 37, 1200, 1999. 66. Rimek, D. et al. Identification of contaminating fungal DNA sequences in zymolyase. J. Clin. Microbiol., 37, 830, 1999. 67. Löffler, J. et al. Comparison of different methods for extraction of DNA of fungal pathogens from cultures and blood. J. Clin. Microbiol., 35, 3311, 1997. 68. Metwally, L. et al. Improving molecular detection of Candida DNA in whole blood: comparison of seven fungal DNA extraction protocols using real-time PCR. J. Med. Microbiol., 57, 296, 2008. 69. Loeffler, J. et al. Automated extraction of genomic DNA from medically important yeast species and filamentous fungi by using the MagNA Pure LC system. J. Clin. Microbiol., 40, 2240, 2002. 70. Sakai. T. et al. A rapid, sensitive and simple detection of Candida deep mycosis by amplification of 18S ribosomal RNA gene: Comparison with assay of serum β-D-glucan level in clinical specimens. Tohoku J. Exp. Med., 190, 119, 2000. 71. Walsh, T.J. et al. PCR and single strand conformational polymorphism for recognition of medically important opportunistic fungi. J. Clin. Microbiol., 33, 3216, 1995. 72. Morace, G. et al. PCR-restriction enzyme analysis for detection of Candida DNA in blood from febrile patients with hematological malignancies. J. Clin. Microbiol., 37, 1871, 1999. 73. Williams, D.W. et al. Identification of Candida species by PCR and restriction fragment length polymorphism analysis of intergenic spacer regions of ribosomal DNA. J. Clin. Microbiol., 33, 2476, 1995. 74. Borst, A. et al. Use of amplified fragment length polymorphism analysis to identify medically important Candida spp., including C. dubliniensis. J. Clin. Microbiol., 41, 1357, 2003. 75. Ahmad, S. et al. Seminested PCR for diagnosis of candidemia: Comparison with culture, antigen detection and biochemical methods for species identification. J. Clin. Microbiol., 40, 2483, 2002. 76. Bagyalakshmi, R., Therese, K.L. and Madhavan, H.N. Application of semi nested polymerase chain reaction targeting internal transcribed spacer region for rapid detection of panfunal genome directly from ocular specimens. Indian J. Opthalmol., 55, 261, 2007. 77. Einsele, H. et al. Detection and identification of fungal pathogens in blood by using molecular probes. J. Clin. Microbiol., 35, 1353, 1997.

563 78. Arancia, S. et al. Construction and use of PCR primers from a 70 kDa heat shock protein gene for identification of Candida albican. Mol. Cell. Probes, 11, 329, 1997. 79. Van Burik, J.A. et al. Panfungal PCR assay for detection of fungal infection in human blood specimens. J. Clin. Microbiol., 36, 1169, 1998. 80. White, P.L., Shetty, A. and Barnes, R.A. Detection of seven Candida species using the Light-Cycler system. J. Med. Microbiol., 52, 229, 2003. 81. Schabereiter-Gurtner, C. et al. Development of novel realtime PCR assays for detection and differentiation of eleven medically important Aspergillus and Candida species in clinical specimens. J. Clin. Microbiol., 45, 906, 2007. 82. McMullan, R. et al. A prospective clinical trial of a real-time polymerase chain reaction assay for the diagnosis of candidemia in nonneutropenic, critically ill adults. Clin. Infect. Dis., 46, 890, 2008. 83. Vollmer, T. et al. Evaluation of novel broad range real-time PCR assay for rapid detection of human pathogenic fungi in various clincial specimens. J. Clin. Microbiol., 46, 1919, 2008. 84. Bašková, L. et al. The Pan-AC assay: a single reaction realtime PCR test for quantitative detection of a broad range of Aspergillus and Candida species. J. Med. Microbiol., 56, 1167, 2007. 85. Innings, A. et al. Multiplex real-time PCR targeting the Rnase P RNA gene for detection and identification of Candida species in blood. J. Clin. Microbiol., 45, 874, 2007. 86. Maaroufi, Y. et al. Rapid detection of Candida albicans in clinical blood samples by using a TaqMan-based PCR assay. J. Clin. Microbiol., 41, 3293, 2003. 87. Guiver, M., Lei, K. and Oppenheim, B.A. Rapid identification of Candida species by TaqMan PCR. J. Clin. Pathol., 54, 362, 2001. 88. Loeffler, J. et al. Quantification of fungal DNA by using fluorescence resonance energy transfer and the light cycler system. J. Clin. Microbiol., 38, 586, 2000. 89. Jordanides, N.E. et al. A prospective study of real-time panfungal PCR for the early diagnosis of invasive fungal infection in haemato-oncology patients. Bone Marrow Transplant, 35, 389, 2005. 90. Klingspor, L. and Jalal, S. Molecular detection and identification of Candida and Aspergillus spp. From clinical samples using real-time PCR. Clin. Microbiol. Infect., 12, 745, 2006. 91. Loeffler, J. et al. Rapid detection of point mutations by fluorescence resonance energy transfer and probe melting curves in Candida species. Clin. Chem., 46, 631, 2000. 92. Widjojoatmodjo, M.N. et al. Nucleic acid sequence-based amplification (NASBA) detection of medically important Candida species. J. Microbiol. Methods, 38, 81, 1999. 93. White, P.L. et al. Detection of Candida in concentrated oral rinse cultures by real-time PCR. J. Clin. Microbiol., 42, 2101, 2000. 94. Tortorano, A.M. et al. Epidemiology of candidaemia in Europe: results of 28 month European confederation of Medical Mycology (ECMM) hospital based surveillance study. Eur. J. Clin. Microbiol. Infect. Dis., 23, 317, 2004. 95. Hope, W., Morton, A. and Eisen, D.P. Increase in prevalence of nosocimial non-Candida albicans candidaemia and the association of Candida krusei with fluconazole use. J. Hosp. Infect., 50, 56, 2002. 96. Colombo, A.L. et al. Brazilian Network Candidemia Study. Epidemiology of candidemia in Brazil: a nationwide sentinel surveillance of candidemia in eleven medical centers. J. Clin. Microbiol., 44, 2816, 2006.

564 97. Al-Tawfiq, J.A. Distribution and epidemiology of Candida species causing fungemia at a Saudi Arabian hospital, 1996– 2004. Int. J. Infect. Dis., 11, 239, 2007. 98. Pfaller, M.A. et al. In vitro susceptibilities of Candida spp to caspofungin: four years of global surveillance. J. Clin. Microbiol., 44, 760, 2006. 99. Uno, K. et al. Evaluation of diagnostic methods for Candida albicans translocation in a mouse model: seminested polymerase chain reaction, blood culture and serological assays. J. Infect. Chemother., 13, 196, 2007. 100. Alam, F.F., Mustafa, A. and Khan, Z.U. Comparative evaluation of (1,3)-β-D-glucan, mannan and anti-mannan antibodies and Candida species specific snPCR in patients with candidemia. BMC Infect. Dis., 7, 103, 2007. 101. Badiee, P. et al. Prospective screening in liver transplant recipients by panfungal PCR-ELISA for early diagnosis of invasive fungal infections. Liver Transplant, 13, 1011, 2007.

Molecular Detection of Foodborne Pathogens 102. Hendolin, P.H. et al. Panfungal PCR and multiplex liquid hybridisation for detection of fungi in tissue specimens. J. Clin. Microbiol., 38, 4186, 2000. 103. Lau, A. et al. Development and clinical application of a panfungal PCR assay to detect and identify fungal DNA in tissue specimens. J. Clin. Microbiol., 45, 380, 2007. 104. Shin, J.H. et al. Rapid detection of up to three Candida species in a single tube by a 5´ exonuclease assay using fluorescent DNA probes. J. Clin. Microbiol., 37, 165, 1999. 105. Löffler, J. et al. Detection of PCR-amplified fungal DNA by using a PCR-ELISA system. Med. Mycol., 36, 275, 1998. 106. Chen, Y.C. et al. Identification of medically important yeasts using PCR-based detection of DNA sequence polymorphisms in the internal transcribed spacer 2 region of the rRNA genes. J. Clin. Microbiol., 38, 2302, 2000. 107. Carvalho, A. et al. Multiplex PCR identification of eight clinically relevant Candida species. Med. Mycol., 45, 619, 2007.

40 Debaryomyces

Juan J. Córdoba, Maria J. Andrade, Elena Bermúdez, Félix Núñez, Miguel A. Asensio, and Mar Rodríguez Universidad de Extremadura

Contents 40.1 Introduction.................................................................................................................................................................... 565 40.1.1 Classification and Morphology of Debaryomyces............................................................................................ 565 40.1.2 Biology, Pathogenesis, and Medical Importance.............................................................................................. 566 40.1.3 Diagnosis of Debaryomyces............................................................................................................................. 567 40.1.3.1 Physiological and Morphological Analysis...................................................................................... 567 40.1.3.2 Molecular Analysis.......................................................................................................................... 568 40.2 Methods.......................................................................................................................................................................... 569 40.2.1 Reagents and Equipment................................................................................................................................... 569 40.2.2 Sample Preparation........................................................................................................................................... 569 40.2.3 Detection Procedures........................................................................................................................................ 570 40.2.3.1 RFLP Analysis of Mitochondrial DNA........................................................................................... 570 40.2.3.2 RAPD-PCR...................................................................................................................................... 571 40.3 Conclusions..................................................................................................................................................................... 571 Acknowledgments...................................................................................................................................................................... 572 References.................................................................................................................................................................................. 572

40.1  Introduction Species of the ascomycetous genus Debaryomyces are among the most common yeasts isolated from many natural habitats such as air, soil, pollen, tree exudates, plants, fruits, insects, feces, gut of vertebrates, and sea water.1 Some of their species, especially Debaryomyces hansenii, have been found in a wide variety of foods, mainly those with low water activity (aw) as well as in high-sugar products such as fruit juices, soft drinks, wine, beer, sugary products, bakery products, dairy products and meat or processed meats.1–12 Also Candida famata, anamorph of D. hansenii, formerly known as Torulopsis candida, can be found in many foods, in particular cheese and other dairy products and sausages.2,13,14 The genus Debaryomyces is normally considered a nonpathogenic yeast15 and has rarely been isolated from humans. Some species have been reported to exert positive effects in the ripening of fermented foods such as dairy and meat products. However, some diseases have been related to D. hansenii and its anamorph C. famata, mainly in immunocompromised patients, such as a case of bone infection,16 an allergic alveolitis,17 or septicemia.18 The positive effects of Debaryomyces in foods and its pathogenic relevance will be discussed in this chapter. For the diagnosis of Debaryomyces, different physiological and morphological methods have been traditionally used. However, these methods are laborious, lack discriminatory power and misidentification occurs frequently.

Progress in the molecular nucleic acid methods may allow the possibility to characterize yeasts at species and strain level. We here will analyze the main physiological and morphological methods, as well as the molecular techniques for the diagnosis of Debaryomyces.

40.1.1 Classification and Morphology of Debaryomyces Genus Debaryomyces was established by Klocker19 with the single species of D. globosus, and currently 18 species have been included: D. carsonii, D. castellii, D. coudertii, D. etchelsii, D. hansenii, D. maramus, D. melissophilus, D. nepalensis, D. occidentalis, D. polymorphus, D. pseudopolymorphus, D. robertsiae, D. udenii, D. vanrijiae, D. yamadae,20 D. prosopidis,21 D. mycophilus,22 and D. singareniensis.23 The members of this genus show spherical cells, and pseudomycelium is absent, primitive or occasionally well developed. All species are perfect, haploid, and have a vegetative reproduction by multilateral budding.24 The sexual reproduction proceeds via heterogamous conjugation of two cells of different form or size, generally mother and bud, although the isogamous conjugation also occurs.24 The conjugation commonly leads to a diplophase followed by meiosis and ascospore formation.25 One to two spherical, globular, ovoidal or lenticular smoothy or warty spores are usually formed per ascus, but in some species up to four spores could be present.24 Debaryomyces species are distinguished from 565

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other ascomycetous yeast genera by the special internal ultrastructure of their ascospores.24 The nuclear base composition of the Debaryomyces species is 37 mol% G + C or higher.26 The kariotype analysis revealed a high degree of polymorphism.27 Thus, in the most frequent species of the genus, D. hansenii, two varieties are differentiated, according to the current taxonomy: D. hansenii var. fabryi and D. hansenii var. hansenii. They can be discriminated by their maximum growth temperatures, the sequence divergences of their 26S rRNA genes, and differences in the electrophoretic mobility of their ­glucose-6-phosphate dehydrogenase.28,29 The anamorph of D. hansenii is C. famata. Phylogene­ tically, D. hansenii is related to Candida albicans30 and belongs to a monophyletic clade containing organisms that translate CTG as serine instead of leucine. For this, Fitzpatrick et al.31 ­suggested that D. hansenii and Candida guilliermondii are sister taxa.

40.1.2 Biology, Pathogenesis, and Medical Importance Debaryomyces species are osmotolerant and can grow in media containing up to 4 M NaCl.32 Species of this genus are characterized physiologically by their inability to assimilate nitrate, as well as their weak or nonexistent fermentation capacities,24 and chemotaxonomically by their expression of coenzyme Q-9.33 These and others characteristics, which are used for testing species of genus Debaryomyces are summarized in Table 40.1. The main species of this genus, D. hansenii is able to grow at 10% NaCl or 5% glucose, and these characteristics are used to discriminate D. hansenii from other ascomycetous yeasts. On the other hand, D. hansenii is one of the

lipid-accumulating, “oleaginous” yeasts, and can accumulate lipids to concentrations up to 70% of its dry biomass34 and its metabolism is clearly dominated by pathways that contribute to lipid metabolism. D. hansenii is a heterogeneous species, with remarkable phenotypic differences between strains, such as variations in their ability to metabolize various carbon sources, the expression of different lipase and protease activities, and their diverse optimal growth conditions.4,35 The biological characteristics of D. hansenii have been reviewed recently by Breuer and Harms.36 D. hansenii can be cultivated in media with up to 25% NaCl or 18% glycerol. It is isolated from environments with high salt concentrations, such as sea water or several types of food. In fact, moderate NaCl concentrations improve growth of D. hansenii cells. The positive effect of NaCl on D. hansenii growth is even more evident in the presence of several stress conditions, such as high temperature and low or high pH levels in the media. All these characteristics make that D. hansenii is regarded as a halophilic yeast.37 D. hansenii is reported to grow optimally at 20–25°C, which might be a consequence of its natural occurrence in habitats such as sea water. D. hansenii can grow at 5°C and even below 0°C. At 10°C this yeast is able to grow at pH 4.0–6.0 when aw is up to 0.99. D. hansenii can grow at aw values as low as 0.65.36,38 The growth of Debaryomyces in foods is not considered usually as harmful, and some species have positive effects in the ripening of fermented foods like dairy and meat products.39–41 Due to their ability to grow at low pH, high salt concentration and low temperature, yeasts are the first microorganisms that develop on the cheese surface. They contribute in cheese ripening by assimilation of lactic acid causing an increase in pH, which will enhance the growth of other microorganisms. In addition, D. hansenii metabolizes lactose,

Table 40.1 Characteristics of Genus Debaryomyces Characteristics

Possible Results for Debaryomyces spp.

Fermentation of: galactose, glucose, lactose, maltose, melibiose, raffinose, sucrose

 + , –, s, v, w

Assimilation of carbon compounds: L-arabinose, cellobiose, citric acid, erythritol, galactose, inositol, lactose, maltose, D-mannitol, raffinose, L-rhamnose, ribitol, D-ribose, soluble starch, succinic acid, sucrose, trehalose, D-xylose Splitting of arbutin

 + , –, v

Assimilation of nitrite

 + , –, v

Growth in vitamin-free medium

 + , –, v  + , –

Growth on 50% (w/w) glucose-yeast extract agar

 + , –, w

Growth at 37°C

 + , –, v 37–40%

G + C (mol%) Shape of the ascospores Wall of the spores Number of spores per ascus Formation of pseudomycelium

spherical, oval warty, smooth 1, 2, 3, 4

 + , – Source: Kreger-van Rij, N.J.W., The Yeast, A taxonomic Study, 3rd ed. Elsevier Science Publishers, Amsterdam, 1984. Notes: +, positive reaction; w, weak reaction; s, slow reaction; –, negative reaction; v, variable reaction.

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as well as multiple organic carbon and nitrogen sources, to generate volatile sulphur compounds, esters, alcohols, aldehydes, and ketones that contribute to alcoholic, acidic, and cheesy flavor.42 In addition, D. hansenii isolated from meat products showed proteolytic and lipolytic activities,43–49 and it has been suggested that D. hansenii enhances the sensory characteristics and may contribute to the flavor in dry-cured fermented sausages,41,50 ripened loins51 or dry-cured hams.12,52 Debaryomyces species also contribute to the ripening of pickles, where they oxidize the acids produced by lactic acid bacteria during fermentation.3,53 It is also possible to find Debaryomyces spp. in other kinds of foods. They were found in ready-to-eat fufu and lafun-fermented cassava products.54 D. polymorphus was one of the most frequently reported yeasts in fruit salads.55 However, D. hansenii has been the most reported species in fermented tea plant (Camellia sinensis) leaves, seasoned green table olives, and processed fresh edible sea urchins.56–58 An excessive growth of Debaryomyces may cause undesirable sensory changes in the formation of bad aromas and flavors, gas production, discoloration, and changes in texture.3,59 Proteolytic D. hansenii was isolated from decayed and damaged, uncooked, ripe tomatoes. Growth of a proteolytic, alkalinizing yeast such as D. hansenii in raw tomatoes enhances conditions for growth of Salmonella, because of increased pH. Thus, the risk of human diseases caused by pathogenic bacteria favored by increased pH of decayed pulp tissue is enhanced by this yeast.60 Debaryomyces spp. have rarely been isolated from humans, but they are considered opportunistic pathogens. Wong et al.16 reported some infections caused by Debaryomyces species. T. candida (C. famata) was isolated from a patient with chronic skin lesions on the hands and feet. D. hansenii was also found in one case of bone infection that a 23-year-old woman suffered over 4 years. Several clinical samples were identified as D. hansenii (and its anamorph C. famata) in superficial infections.61 D. hansenii was also responsible for a persistent candidemia observed in a patient heavily treated with various fungicides.18 A 65-year-old female was diagnosed of extrinsic allergic alveolitis resulting from exposure to inhaled organic dusts, being D. hansenii the dominant species in indoor air samplings.17 Different species of Candida are part of the natural microbiota and, thus, are regarded as commensal organisms in humans. C. famata was thought to be nonpathogenic for humans. However, this yeast was isolated in combination with other Candida spp. from a relevant number of clinical cases, including ocular endophthalmitis, retinopathy, and central nervous system infection. In addition, C. famata is rarely implicated in human fungemia.62–64 New treatments of hospitalized patients seem to have favored the emergence of C. famata as a pathogen, but they may be ­under-reported.65 In this sense, C. famata has been associated with ­catheter-related bloodstream infection66 and rarely with other infections,16,67–71 generally in immunocompromised patients. Thus, medical

importance of D. hansenii and its anamorph C. famata may rely on susceptibility of immunocompromised patients and on its resistance to the treatments applied to different pathologies.

40.1.3  Diagnosis of Debaryomyces Different physiological and morphological methods have been traditionally used in taxonomic differentiation of Debaryomyces.1,33,72–74 However, several studies have shown that traditional identification methods, based on phenotypic properties of yeasts (morphological, biochemical, and physiological tests) are laborious, lack discriminatory power and misidentification occurs frequently.75 In addition, identification on the basis of morphological properties generates the so-called double binomial nomenclature, with one name for the vegetative state (anamorph) and another for the sexual state (teleomorph). This is the case of C. famata/D. hansenii. Furthermore, these methods generally produce ambiguities and inaccuracies in the results, because the morphological and physiological characteristics are strongly influenced by growing conditions. Progress in molecular biology in the last decade has opened up possibilities of characterizing yeasts at the genomic level. The sequencing of the genes coding for 18S and 26S ribosomal RNA (rRNA), as well as internal transcribed spacer (ITS), has brought about many changes in the identification and classification of yeasts.76 In addition, techniques based on random amplified polymorphic DNA (RAPD-PCR) and restriction fragment length polymorphism (RFLP), have already been recognized as reliable tools for the rapid identification of yeasts.9 We here review the main physiological and morphological methods, as well as the molecular techniques. 40.1.3.1  Physiological and Morphological Analysis The physiological and morphological identification of Debaryomyces is done on the basis of several characteristics of this genus which are listed below: Characteristics of vegetative reproduction. (i) Modes of vegetative reproduction: by budding, by fission or a combination of both processes; (ii) characteristic of vegetative cells: morphology grown in liquid and solid media; (iii) formation of pseudomycelium and true mycelium; (iv) formation of asexual endospores; (v) formation of chlamydospores; and (vi) formation of germ tubes. Sexual characteristics. Characteristics of ascospore or basidiospore formation. Physiological and biochemical characteristics. The physiological tests used for identifying purposes are those associated with the utilization of carbon and nitrogen sources, growth factor requirements, growth at elevated temperatures and on media of high sugar or sodium chloride content, formation of typical characteristic metabolites, and susceptibility to antibiotics. The utilization of these tests for Debaryomyces characterization requires considerable experience and skill for evaluating specified

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Molecular Detection of Foodborne Pathogens

tests. Furthermore, great difficulties for the differentiation at species level in this genus could be found, since many biochemical and physiological tests show the same result for different species. Among the physiological biochemical tests of characterization, in addition to those in Table 40.1, the following have been used: • • • • • • • • • •

Formation of extracellular, amyloid compounds Production of ammonia from urea Splitting of fat Ester production Cycloheximide resistance Tolerance of 1% acetic acid Gelatin liquefaction Diazonium Blue B (DBB) color test Coenzyme Q structure Tetrazolium indicator medium (TTC medium)

The main biochemical tests are electrophoresis of proteins, coenzyme Q analysis, allozyme analysis, and ultrastructure and chemical composition (polysaccharides, fatty acids) of the cell wall.24 Missoni et al.77 found the evaluation of cell fatty acids by gas chromatography very useful in the routine diagnosis and epidemiological monitoring of the infection due to Candida spp. (including C. famata). These techniques are not always stable or reproducible because they depend on the physiological status of the strains. For example, fermentation of sugars is not very accurate because the slow release of CO2 is not so immediate to be trapped in a tube Durham. Many commercial methods based on the above morphological and physiological characteristics have been developed for the identification of yeasts, although most of them focus on clinical isolates related to different diseases that do not include species of genus Debaryomyces. Some examples of this are the Yeast Identification System API 20C (BioMérieux), the UniYeast-Yek system (Remel), the Minitek system (BBL),72,78–80 or enzyme-based system such as Yeast Identification Panel (Baxter-MicroScan) and MicroScan Rapid Yeast Identification (Innovative Diagnostic Systems).73,81 The basic principle of these systems is carbohydrate assimilation, which requires a minimum incubation period of 24 h for growth. The enzymatic activity systems use chromogenic substrates and can identify yeasts within 4 h after inoculation. Due to the increasing importance of yeasts in the production and spoilage of foods, the database of the above methods has been completed with foodborne yeasts. Thus, the API ID32C system (BioMérieux) allows the identification of three species of the genus Debaryomyces: D. hansenii, D. marama, and D. polymorphus.82 Similarly, the Vitek Yeast Biochemical card (BioMérieux) allows identification of D. hansenii.83,84 Furthermore, several methods have been developed for automatic identification of yeasts on the basis of biochemical tests, such as the highlighting system Vitek 2® (BioMérieux) with colorimetric and fluorimetric VITEK 2 yeast cards85,86 and Biolog YT Microplate® (AES Laboratories).

In addition, selective, and differential chromogenic solid media have been developed for the detection of Debaryomyces spp. For food samples, Debaryomyces differential medium (DDM) has been reported to be very satisfactory.87,88 40.1.3.2  Molecular Analysis The molecular methods for identifying genus Debaryomyces and yeasts in general, are based on the study of DNA and RNA. The yeast nucleic acid sequences contain the primary information which determines all the physiological, biotechnological, and pathogenic characteristics and potential of a particular organism. The use of genotypic rather than phenotypic characteristics for identification is potentially more accurate, reproducible, and rapid.83 Nucleic acid-based methods have the advantage over phenotypic identification methods by not being influenced by environmental conditions of the cells, because the nucleotide sequence of the DNA does not change during growth. (i) Methods based on nucleic acid hybridization: Nucleic acid hybridization is typically between a DNA or RNA molecule present in the target organism and a DNA probe which has a sequence complementary to the target sequence. 18S rRNA-targeted oligonucleotide probes were designed for rapid and reliable identification of yeasts like the genus Debaryomyces and the species D. hansenii.89–91 (ii) Methods based on nucleic acid amplification: The most popular method of amplification is the polymerase chain reaction (PCR) technique. The PCR can detect one copy of the target sequence by using two oligonucleotide primers. Conventional PCR has been used for identification of D. hansenii only in some cases.92 However, different variations of this technique (PCR-RFLP, RAPD, Q-PCR, NASBA, etc.) are frequently used to differentiate the genus Debaryomyces. One of the problems when using PCR with clinical samples or isolates related to pathogenesis is the possibility of detecting naked DNA derived from dead and degrading yeast cells instead of live yeasts, which results in false-positives. An alternative to PCR consists in using nucleic acid sequencebased amplification (NASBA) system that selectively amplifies RNA. This method has been used with yeasts of genus Candida.93 The RAPD-PCR is a variation of PCR used for the identification of yeasts. This method is based on PCR amplification of the genomic DNA in the presence of a single short primer. Due to low-temperature hybridization primer joins unspecific sites throughout the genome, allowing the amplification of DNA fragment of different length. The use of RAPD-PCR permits to obtain fingerprints which are specific for species and even strains. The RAPD-PCR technology with different primers, mainly from micro- and minisatellites, has been used for the correct identification of several species of Debaryomyces.75,83,94–96

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One of the most promising PCR techniques in the detection of microorganisms is real-time PCR or quantitative PCR (Q-PCR). Q-PCR assays have been developed for detecting and enumerating yeasts, specially for yeasts in wine97,98 and foods,99 as well as in fungal infections by C. famata.64

The most reliable, simple, and fast methods to differentiate Debaryomyces spp. at strain level rely first on RFLP analysis of mtDNA and then on RAPD-PCR using micro- or minisatellite primers.94

(iii) Methods based on RFLP: The determination of the RFLP is the differentiation of organisms by analyzing patterns of rupture that are generated in a specific site of the genome when it is cut by restriction enzymes. Then, gel electrophoresis displays a pattern of polymorphic bands corresponding to the fragments of different sizes, which are generated on the cut of each endonuclease. This fragment length polymorphism appears because the organisms of different species and even strains differ in the distance of cleavage sites for each restriction enzyme. The similarity of the patterns generated allows for correlations between species and strains. The PCR-RFLP is a useful method for identification of some yeast genus, using the restriction analysis of different regions of ribosomal RNA/DNA genes (ITS, 5.8S rDNA, 18S rDNA, etc.). This technique has two steps: first, the rDNA region is amplified and then the PCR product is digested. Therefore, this technique is slower than some current PCR methods, which achieve identification and typing with just the PCR products. The results of this technique depend on the region of genome amplified and the enzymes used. For example, PCR-RFLP of ITS1-5.8S rDNA-ITS2 and 18S rDNA regions has been reported as a good method for the differentiation of Saccharomyces species,100,101 but it does not allow for separation of Debaryomyces spp.94 However, the PCR-RFLP of the intragenic spacer (IGS) of rDNA is proposed as a clear technique for the practical discrimination species of the genus Debaryomyces.102 In the technique of pulsed-field gel electrophoresis (PFGE), restriction enzymes digest the complete genome and large DNA molecules are resolved by continuous reorientation of the electric field during gel electrophoresis, determining chromosome length polymorphism (CLP). This pattern is specific for yeast species, due to genetic and evolutionary phenomena that have taken place in the chromosomes (insertions, deletions, and translocations). This method has been reported as a useful tool in the differentiation of species and strains of Debaryomyces.27,35 Another technique based on RFLP is the mitochondrial DNA (mtDNA) restriction analysis. Among all the molecular techniques described in literature, mtDNA restriction analysis appears as one of the most suitable methods to differentiate between yeast strains. Querol et al.103 developed a new mitochondrial restriction analysis method based on the extraction of total yeast DNA and the use of GC-rich restriction endonucleases that recognize a high number of sites in the yeast nuclear DNA, but few sites in the mtDNA. This technique has successfully been used to characterize strains of genus Debaryomyces.94,104–106

40.2  Methods In this chapter, a differentiation method for Debaryomyces spp. that includes first mtDNA restriction analysis and then RAPD-PCR using micro- and minisatellite primers is described. For this, total yeast DNA is first isolated and then either digested with HaeIII for RFLP of mtDNA or amplified by RAPD-PCR using microsatellite primers (GACA)4, (GAC)5 and (GTG)5 and the minisatellite primer M13.

40.2.1 Reagents and Equipment Reagents and equipment for RAPD-PCR and mtDNA restriction analysis are listed in Table 40.2.

40.2.2 Sample Preparation Samples must be taken aseptically and homogenized in a Stomacher lab-blender with sterile peptone water (0.1%, w/v). Decimal dilutions are obtained with the same diluent and 0.1 ml is spread onto the surface of different selective media for yeasts, such as Dichloran Rose-Bengal Chloramphenicol Agar (DRBC, Oxoid, Cambridge, UK), Malt Extract Agar (2% w/v malt extract, 2% w/v glucose, 0.1% w/v peptone, 2% w/v agar) and Dichloran-Glycerol Agar (DG18, Oxoid). They are incubated at 25°C for 5 days. The isolates obtained can be placed in cryotubes containing Malt Extract Broth (2% w/v malt extract, 2% w/v glucose, 0.1% w/v peptone) and stored at –80°C in 20% (v/v) glycerol. Total DNA is isolated from broth cultures. Yeast cells are grown in 10 ml of Yeast Peptone Glucose broth (1% w/v yeast extract; 2% w/v peptone; 2% w/v glucose) in a 50 ml conical tube at 25°C in an orbital shaker at 200 rpm (Figure 40.1). In addition, for mitochondrial DNA isolation, conical tubes should be placed inclined in the orbital shaker. To reach an adequate cell density, incubation for 48 h is recommended. Cells are pelleted by centrifugation for 5 min at 4,000 rpm, resuspended in 500 µl of 1 M sorbitol, 0.1 M EDTA, pH 7.5, and transferred to a 2 ml sterile microtube. Subsequently, 40 µl of lyticase (20 mg/ml) are added to digest cell walls of yeasts and obtain spheroplasts. After 45 min of incubation at 37°C in a water bath with occasional shaking, the suspension is centrifuged at 13,000 rpm for 1 min and the supernatant is discarded. The pellet is resuspended in 500 µl of 50 mM Tris-HCl, 20 mM EDTA pH 7.4 to release cellular DNA from spheroplasts. After that, 50 µl of 10% (w/v) sodium dodecyl sulfate (SDS) are added and the mixture is heated at 65°C for 10 min in a water bath. To remove proteins, 200 µl of 5 M potassium acetate are added, the solution is shaken and stored on ice for 15 min. Next, it is centrifuged at 13,000 rpm

570

Molecular Detection of Foodborne Pathogens

Table 40.2 Reagents and Equipment Required for RFLP Analysis of Mitochondrial DNA and RAPD-PCR Reagents and Culture Media Glycerol

Isopropanol

Lyticase

Ethanol absolute

Sodium dodecyl sulfate (SDS), molecular biology grade

Ethyllendiaminetetraacetic acid (EDTA) disodium salt, dihydrate

Yeastcells 10 ml YPG broth Incubation Centrifugation Resuspension Cell wall digestion

D-Sorbitol extrapure

HaeIII and 10 × buffer

Hydrochloric Acid (HCl) 35%

Agarose D-1

Spheroplasts

Potassium acetate, extra pure

DNA molecular markers

Centrifugation

RNase

Bromophenol blue, indicator

Ultrapure water

Ethidium bromide

PCR reagents (see Table 40.3)

Bacteriological agar

Dichloran 18% Glycerol (DG18) Agar

Dichloran Rose Bengal Chloramphenicol (DRBC) Agar D-Glucose anhydrous, extra pure Bacteriological peptone

Malt extract Yeast extract Tris-(hydroxymethyl)aminomethane

Resuspension Protein resolubilization Protein precipitation Centrifugation DNA precipitation Centrifugation

Centrifugation

Laminar flow cabinet

Incubator

Stomacher lab-blender

Orbital shaker

Vortex

Freezer

Water bath

Spectrophotometer

RNA digestion

Thermal cycler and PCR tubes

5 ml cryotubes

IsolatedDNA

Centrifuge for 2 ml and 0.5 ml microtubes

1 ml, 200 µl, 50 µl, 5 µl, 1 µl and 0.5 µl pipettes UV transilluminator

Microwave oven

(4,000 rpm, 5 min) Supernatant (500 µl 1 M sorbitol, 0.1 M EDTA, pH 7.5) (40 µl lyticase 20 mg/ml, 37°C, 45 min)

(13,000 rpm, 1min) Supernatant (500µl 50mM Tris-HCl, 20m MEDTA, pH7.4)

Free DNA

Whasing Equipment

(25 °C, 200 rpm, 48 h)

Drying Dissolution

(50 µl 10% SDS, 65°C, 10 min) (200 µl 5 M KOAc, 0°C, 15 min) (13,000 rpm, 5 min) Pellet (500 µl isopropanol, 5 min) (13,000 rpm, 10 min) Supernatant (1 ml 70% ethanol) (13,000 rpm, 1 min) Supernatant (37°C, 45 min) (50 µl TE buffer) (1 µl 20 mg/ml RNase, 37°C, 30 min) (–20 °C)

Figure 40.1  Diagram of DNA isolation from Debaryomyces yeasts.

Horizontal electrophoresis unit with Centrifuge for 50 ml conical tubes the appropriate gel casting tray and combs

40.2.3  Detection Procedures

for 5 min. A volume of 500 µl of the resulting supernatant is transferred to a new sterile microtube together with an equal volume of ice-cold isopropanol and it is left at room temperature for 5 min. DNA is pelleted by centrifugation at 13,000 rpm for 10 min, washed with 1 ml of ice-cold 70% (v/v) ethanol and centrifuged at 13,000 rpm for 1 min. Ethanol is aspirated with a pipette and the pellet is dried at 37°C for 45 min. Dried DNA is dissolved in 50 µl of TE buffer (10 mM Tris-HCl, 1mM EDTA pH 8.0). For RNA digestion, 1 µl of RNase (20 mg/ml) is added and the solution is incubated at 37°C for 30 min in a water bath and immediately placed at –20°C. The DNA obtained can be used for RFLP analysis and RAPD-PCR. To be used for PCR the DNA has to be spectrophotometrically quantified and brought to a final concentration of 100 ng/µl.

40.2.3.1 RFLP Analysis of Mitochondrial DNA Mitochondrial DNA restriction analysis consists in the digestion of total DNA with restriction endonucleases with recognition sites rich in GC, such as HaeIII, that results in an overdigestion of the nuclear DNA to render specific bands from mtDNA. The digestion mixture is prepared on ice for a final volume of 15 µl as follows: 5 µl of DNA isolated according to the above method, 20 U of the restriction enzyme HaeIII and 1.5 µl of the appropriate 10 × buffer.94 Then, the reaction is performed overnight at 37°C in a water bath (Figure 40.2). An alternative method using a microwave oven for DNA digestion has been described.107,108 The digestion mixture is placed inside a water bath and incubated in three heating times at maximum level of the microwave oven (1,250 W) for 20 s each, giving a spin between each time.

571

Debaryomyces

Isolated DNA Digestion mixture

(5 µl DNA, 20 U Hae III, 1.5 µl 10 × buffer) Microwave oven (1,250 W, 3 × 20 s)

Water bath (37ºC, overnight) Digested DNA Sample preparation

(molecular marker, loading buffer) Electrophoresis gel (0.8% agarose in TAE buffer) – +

Electrophoresis

Visualization

(100 V)

(0.5 µg/ml ethidium bromide, UV)

Figure 40.2  Diagram of RFLP analysis of mitochondrial DNA from Debaryomyces yeasts.

Restriction fragments are electrophoretically separated in a horizontal 0.8% (w/v) agarose gel in 1 × TAE buffer (40 mM Tris-acetate, 1 mM EDTA pH 8.0) at 100 V. Each digested DNA sample, as well as an appropriate DNA molecular marker, are mixed with loading buffer (50% glycerol, 0.25% bromophenol blue and 25 mM EDTA) and loaded into the gel wells. Restriction fragments are visualized in an UV transilluminator after ethidium bromide (0.5 µg/ml) staining. Sizes of restriction fragments can be estimated by comparison to the DNA molecular marker. 40.2.3.2 RAPD-PCR For RAPD-PCR, a single primer with an arbitrary sequence of oligonucleotides is used. Thus, knowledge of DNA template sequence from the tested yeasts is not required. Core sequence of phage M13 (5´-GAGGGTGGCGGTTCT-3´)109 and the microsatellite primers (GACA)4, (GAC)5 and (GTG)5 have proved useful to characterize Debaryomyces spp. using RAPD-PCR.94 The PCR mixture must be prepared on ice with the reagents and concentrations summarized in Table 40.3 to reach a final volume of 50 µl. The reaction is performed in a thermal cycler following the amplification programs shown in Table 40.3. After the program ends, RAPD-PCR products are kept in the thermal cycler at 4°C. The reaction mixture adding water instead of DNA sample can be used as negative control. RAPD-PCR products are examined by electrophoresis in horizontal 1% (w/v) agarose gels at 100 V (Figure 40.3).

A DNA molecular marker can be incorporated into the gel to estimate the size of the amplification products. Gels are stained with an ethidium bromide (0.5 µg/ml) solution and visualized under UV light.

40.3  Conclusions Yeasts of genus Debaryomyces have been usually found as microbial population of food, specially ripened foods. These yeasts are considered normally as nonpathogenic. However, some species of this genus have been occasionally isolated from human diseases such as bone infection, allergic alveolitis or septicaemia in immunocompromised patients. Eighteen different species have been included in the genus Debaryomyces. All of them are osmotolerant and can grow in media containing up to 4 M NaCl. Species of this genus are characterized physiologically by their inability to assimilate nitrate, as well as their weak or nonexistent fermentation capacities. Among the physiological, morphological, and molecular methods used for characterizing Debaryomyces species, the most reliable, simple, and fast way to characterize Debaryomyces spp. relies on RFLP analysis of mtDNA and RAPD-PCR using micro- or minisatellite primers. In the present work, a combined procedure that includes characterization by both above methods to differentiate Debaryomyces spp. at strain level has been described.

572

Molecular Detection of Foodborne Pathogens

Table 40.3 RAPD-PCR Reagents and Programs for Different Primers M13 Reagents

Stock Concentration 10 mM Tris-HCl (pH 8.8), 50 mM KCl, 0.1% Triton X-100

Mg2 + -free reaction buffer

(GACA)4

(GAC)5

(GTG)5

Volume (µl) 5

5

5

5

MgCl2

50 mM

2

3

4

4

PCR nucleotide mix

10 mM

1

1

1

1

Primer

100 ng/µl

1

2

2

2

DNA

100 ng/µl

2

1

1

1

Taq DNA polymerase

2 U/µl

0.5

0.5

0.5

0.5

Sterile deionized water

 

38.5

37.5

36.5

36.5

PCR stages

Number of Cycles

Initial denaturation

PCR Program

1

94°C, 3 min

 

94°C, 45 s

94°C, 1 min

94°C, 30 s

30

50°C, 1 min

36°C, 1 min

45°C, 1 min

60°C, 3 min

55°C, 5 min

60°C, 5 min

1

60°C, 3 min

55°C, 5 min

60°C, 5 min

Denaturation Annealing Extension Final extension

Isolated DNA

5’GAGGGTGGCGGTTCT3’ (GACA)4, (GAC)5, (GTG)5

Primers

Sample preparation

(Reaction buffer, MgCl2, nucleotide mix, Taq polymerase, water)

Amplification

Sample preparation

94°C, 5 min

(See Table 3)

(Molecular marker, loading buffer) Electrophoresis gel (1% agarose in TAE buffer)

+ Electrophoresis

Visualization

– (100 V)

(0.5 µg/ml ethidium bromide, UV)

Figure 40.3  Diagram of RAPD-PCR from Debaryomyces yeasts.

Acknowledgments

References

This work was funded by part of the projects AGL2004-03291 and AGL2007-64639 funded by the Spanish Ministerio de Educación y Ciencia.



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574 45. Martín, A. et al. Contribution of a selected fungal population to proteolysis on dry-cured ham. Int. J. Food Microbiol., 94, 55, 2004. 46. Bolumar T. et al. Purification and characterization of a prolyl aminopeptidase from Debaryomyces hansenii. Appl. Environ. Microbiol., 69, 227, 2003. 47. Bolumar T. et al. Purification and properties of an arginyl aminopeptidase from Debaryomyces hansenii. Int. J. Food Microbiol., 86, 141, 2003. 48. Bolumar, T. et al. Protease B from Debaryomyces hansenii: purification and biochemical properties. Int. J. Food Microbiol., 98, 167, 2005. 49. Durá, M.A., Flores, M. and Toldrá, F. Effect of Debaryomyces spp. on the proteolysis of dry-fermented sausages. Meat Sci., 68, 319, 2004. 50. Flores, M., et al. Effect of Debaryomyces spp. on aroma formation and sensory quality of dry-fermented sausages. Meat Sci., 68, 439, 2004. 51. Martín, A. et al. Effect of Penicillium chrysogenum and Debaryomyces hansenii on the volatile compounds during controlled ripening of pork loins. Int. J. Food Microbiol., 84, 327, 2003. 52. Martín, A. et al. Contribution of a selected fungal population to the volatile compounds on dry-cured ham. Int. J. Food Microbiol., 110, 8, 2006. 53. Boekhout, T. and Phaff, J. Yeast biodiversity. In: Yeasts in Food, Beneficial and Detrimental Aspects, Boekhout, T. and Robert, V., Eds. Behr’s Verslag, Hamburg, 2003. 54. Omafuvbe, B.O. et al. Microbial diversity in ready-to-eat fufu and lafun-fermented cassava products sold in Ile-Ife, Nigeria. Res. J. Microbiol., 2, 831, 2007. 55. Tournas, V.H., Heeres, J. and Burgess, L. Moulds and yeasts in fruit salads and fruit juices. Food Microbiol., 23, 684, 2006. 56. Kajikazawa, T. et al. Detection of pathogenic yeasts from processed fresh edible sea urchins sold in a fish market. Jpn. J. Med. Mycol., 48, 169, 2007. 57. Sansone, C. et al. Isolation of a psychrotolerant Debaryomyces hansenii strain from fermented tea plant (Camellia sinensis) leaves. J. Plant Interactions, 2, 169, 2007. 58. Hernández, A. et al. Determination of killer activity in yeasts isolated from the elaboration of seasoned green table olives. Int. J. Food Microbiol., 121, 178, 2008. 59. Jakobsen, M. and Narvhus, J. Yeasts and their possible beneficial and negative effects on the quality of dairy products. Int. Dairy J., 6, 755, 1996. 60. Wade, W.N. et al. Proteolytic yeasts isolated from raw, ripe tomatoes and metabiotic association of Geotrichum candidum with Salmonella. Int. J. Food Microbiol., 86, 101, 2003. 61. Nishikawa, A. et al. Taxonomic position of clinical isolates of Candida famata. J. Med. Vet. Mycol., 34, 411, 1996. 62. Krcmery, V. and Barnes, A.J. Non-albicans Candida spp. causing fungaemia: pathogenicity and antifungal resistance. J. Hospital Infect., 50, 243, 2002. 63. Carrasco, L. et al. Isolation of Candida famata from a patient with acute zonal occult outer retinopathy. J. Clin. Microbiol., 43, 635, 2005. 64. Pisa, D. et al. Evolution of antibody response and fungal antigens in the serum of a patient infected with Candida famata. J. Med. Microbiol., 56, 571, 2007. 65. Hazen, K.C. New and emerging yeast pathogens. Clin. Microbiol. Rev., 8, 462, 1995. 66. Ng, K.P. et al. Scopulariopsis brevicaulis infection in a patient with acute myeloid leukemia. Med. J. Malaysia, 58, 608, 2003.

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Debaryomyces 87. De Silóniz, M.I., Valderrama, M.J. and Peinado, J.M. A chromogenic medium for the detection of yeasts with beta galactosidase and beta-glucuronidase activities from intermediate moisture foods. J. Food Prot., 68, 808, 2000. 88. Quirós, M. et al. A beta-glucuronidase-based agar medium for the differential detection of the yeast Debaryomyces hansenii from foods J. Food Prot., 68, 808, 2006. 89. Kosse, D. et al. Identification of yoghurt-spoiling yeasts with 18S rRNA-targeted oligonucleotide probes. Syst. Appl. Microbiol., 20, 468, 1997. 90. Corredor, M. et al. DNA probes specific for the species Debaryomyces hansenii: useful tools for rapid identification. FEMS Microbiol. Lett., 193, 171, 2000. 91. Davila, A.M. et al. Use of specific DNA probes for the rapid characterization of yeasts isolated from complex biotypes. Gen. Sel. Evol., 33, 353, 2001. 92. Nishikawa, A., Sugita, T. and Shinoda, T. Rapid identification of Debaryomyces hansenii/Candida famata by polymerase chain reaction. Med. Mycology, 37, 101, 1999. 93. Borst, A. et al. Detection of Candida sp in blood cultures using nucleic acid sequence-based amplification (NASBA). Diag. Microbiol. Inf. Dis., 39, 155, 2001. 94. Andrade, M.J. et al. DNA typing methods for differentiation of yeasts related to dry-cured meat products. Int. J. Food Microbiol., 107, 48, 2006. 95. Baruzzi, F. et al. Molecular and physiological characterization of natural microbial communities isolated from a traditional Southern Italian processed sausage. Meat Sci., 72, 261, 2006. 96. Senses-Ergul, S. et al. Characterization of some yeasts isolated from foods by traditional and molecular tests. Int. J. Food Microbiol., 108, 120, 2006. 97. Hierro, N. et al. Real-time quantitative PCR (QPCR) and reverse transcription-QPCR for detection and enumeration of total yeasts in wine. Appl. Environ. Microbiol., 72, 7148, 2006. 98. Phister, T.G. and Mills, D.A. Real-time PCR assays for detection and enumeration of Dekkera bruxellensis in wine. Appl. Environ. Microbiol., 69, 7430, 2003.

575 99. Larpin, S. et al. Geotrichum candidum dominates in yeast population dynamics in Livarot, a French red-smear cheese. FEMS Yeast Res., 6, 1243, 2006. 100. Guillamón, J.M. et al. Rapid identification of wine yeast species based on RFLP analysis of the ribosomal internal transcribed spacer (ITS) region. Arch. Microbiol., 169, 387, 1997. 101. Jespersen, L., Kühle, A. and Petersen, K.M. Phenotypic and genetic diversity of Saccharomyces contaminants isolated from lager breweries and their phylogenetic relationship with brewing yeasts. Int. J. Food Microbiol., 60, 43, 2000. 102. Quirós, M. et al. PCR-RFLP analysis of the IGS region of rDNA: a useful tool for the practical discrimination between species of the genus Debaryomyces. Ant. Leeuwenhoek, 90, 211, 2006. 103. Querol, A., Barrio, E. and Ramón, D. A comparative study of different methods of yeast strain characterization. Syst. Appl. Microbiol., 15, 439, 1992. 104. Romano, A. et al. Use a RAPD and mitochondrial DNA RFLP for typing of Candida zeylanoides and Debaryomyces hansenii yeast strains isolated from cheese. Syst. Appl. Microbiol., 19, 255, 1996. 105. Petersen, K.M., Moller, P.L. and Jespersen, L. DNA typing methods for differentiation of Debaryomyces hansenii strains and other yeasts related to surface ripening cheeses. Int. J. Food Microbiol., 69, 11, 2001. 106. Martorell, P., Fernández-Espinar, M.T. and Querol, A. Molecular monitoring of spoilage yeasts during the production of candied fruit nougats to determined food contamination sources. Int. J. Food Microbiol., 101, 293, 2005. 107. López, V. et al. A simplified procedure to analyse mitochondrial DNA from industrial yeasts. Int. J. Food Microbiol., 68, 75, 2001. 108. Casado, E.M. et al. Rapid Characterization of yeasts isolated from Iberian ham by restriction analysis Mitochondrial DNA. XX National Congress of Microbiology, Caceres, 2005. 109. Huey, B. and Hall, J. Hypervariable DNA fingerprinting in Escherichia coli. Minisatellite probe from bacteriophage M13. J. Bacteriol., 171, 2528, 1989.

41 Fusarium

Antonio Moretti and Antonia Susca Institute of Sciences of Food Production

Contents 41.1 Introduction.................................................................................................................................................................... 577 41.1.1 Classification and Morphology of Major Foodborne Fusarium....................................................................... 577 41.1.2 Biology and Pathogenesis................................................................................................................................. 579 41.1.3 Diagnosis of Fusarium..................................................................................................................................... 581 41.1.3.1 Fumonisins-Producing Species........................................................................................................ 582 41.1.3.2 Trichothecenes Producing Species.................................................................................................. 583 41.1.3.3 Surface Plasmon Resonance (SPR) Biosensors............................................................................... 585 41.2 Methods.......................................................................................................................................................................... 585 41.2.1 Molecular Detection of Fumonisins-Producing Fusarium spp. in Maize........................................................ 585 41.2.2 Quantitative Detection of Fumonisins-Producing Fusarium spp. in Maize..................................................... 586 41.2.3 Molecular Detection of Trichothecenes Fusarium spp. in Wheat.................................................................... 586 41.3 Conclusions and Future Perspectives............................................................................................................................. 587 References.................................................................................................................................................................................. 588

41.1  Introduction The genus Fusarium comprises a high number of fungal species that can be plant-pathogenic, causing diseases in several agriculturally important crops including cereals, and also can be harmful for humans and animals. Many of them produce a wide range of biologically active secondary metabolites (e.g., mycotoxins) with an extraordinary chemical diversity. The biological activity of Fusarium mycotoxins can be detrimental to plants, and associated with cancer and other diseases in humans and domesticated animals. The combined effect of Fusarium species infecting several crops and producing mycotoxins in the field is the contamination of cereal grains and other plant-based foods. With many pathogenic and opportunistic species of the genus colonizing plants as part of a complex of Fusarium species, it provides an interesting example of biodiversity as well as a consequence of the different environmental conditions that exist in the various agro-ecosystems in which crops are cultivated. These conditions can also influence the fungal-plant interactions of the single species and their capability to produce mycotoxins. Moreover, the ability of various Fusarium species within the complexes to produce different classes of secondary metabolites combined with their ability to coexist in the same host or/and occur in quick succession have allowed these complexes to become “invincible armadas” against many plants.1 Plant infections by Fusarium can occur at all developmental stages, from germinating seeds to mature vegetative tissues, depending on the host plant and Fusarium species involved. Therefore, since most Fusarium species have specific mycotoxin profiles, early, and accurate identification of the Fusarium species occurring in the plants at every step of

their growth is critical to predict the potential toxicological risk to which plants are exposed and to prevent toxins entering the food chain. However, the unambiguous identification of mycotoxigenic Fusarium species still remains a most critical issue, given that the number of species (which stands now over 80)2 recognized in the genus has been constantly changing in the last century in accordance with the different taxonomic systems. Furthermore, this genus provides few morphological characters useful for species discrimination based only on traditional techniques, although, fortunately, some of the most important toxigenic and pathogenic Fusarium species can be diagnosed, with some experience, by using only their morphological traits. Considering that the current criteria for Fusarium species identification [e.g., morphological (MSR),3,4 biological (BSR)5 and phylogenetic species recognition (PSR)6] rarely agree, and that there are over 81 of the 101 most economically important plants each having at least one associated Fusarium disease2 and each Fusarium species keeping its own toxicological profile,7 it is a challenge to ascertain the taxonomic status of Fusarium species on their phenotypical characteristics (including pathogenicity and toxigenicity) alone (www.apsnet.org/online/ common/search.asp).

41.1.1 Classification and Morphology of Major Foodborne Fusarium The genus Fusarium belongs to the Ascomycota phylum, Ascomycetes class, Hypocreales order,8 while the teleomorphs of Fusarium species are mostly classified in the genus Gibberella and for a smaller number of species in the 577

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Hemanectria and Albonectria genera. For a complete review of the main taxonomic systems that have contributed to define the modern taxonomy of Fusarium, see the excellent work of Leslie and Summerell,2 which contains an updated description, not just morphological, of 70 species within the genus. The main approach for the Fusarium classification is still the morphology and the primary trait for species to be placed in Fusarium genus is the occurrence of the asexual spores, the distinctive banana-shaped macroconidia, firstly diagnosed by Link.9 Fusarium species produce three types of spores: macroconidia, microconidia, and chlamydospores.2 Septated macroconidia can be produced on monophialides and polyphialides in the aerial mycelium, but often also on short monophialides in specialized structures called sporodochia.10 A monophialide is a conidiation cell with a unique pore from which the endoconidia are released; a polyphialide can possess several such openings. Microconidia can vary in shape and size, and are produced in the aerial mycelium in clumps or chains, both on monophialides and polyphialides. Finally, chlamydospores are resistance structures with thickened walls and a high lipid content; when they are present, they can form in the middle of the hyphae or at their termini. The different shape of macroconidia remains the most important character for distinguishing the species. Moreover, other traits such as the presence/absence of microconidia and their shape, the presence/absence of chlamydospores, and the characters of the micro- and macro-conidiogenous cells contribute to distinguish species in Fusarium. In order to identify the species, all taxonomists suggest to use strain cultures deriving from single-spore isolation and grow the strains on special media under standard incubation conditions. All taxonomic systems developed so far are based on a seminal work by Wollenweber and Reinking11 with various modifications.4,12 This publication organized the genus in 16 Sections including 65 species, 55 varieties and 22 forms. The main discriminating criteria among Sections were based on morphology; in particular on the presence and shape of microconidia, on the presence and position of chlamydospores in the hyphae, on the shape of macroconidia and of their basal cells. The taxonomic system described by Gerlach and Nirenberg3 kept the number of Sections as Wollenweber and Reinking, while Nelson et al.4 proposed a simpler classification method that divided the genus in 12 Sections. Although the main taxonomic systems have organized Sections with species sharing common morphological characters and thus supposedly genetically related, not all researchers accept Section concept since some of the morphological characters used are now considered of poor reliability from an evolutionary point of view, according with recent molecular investigations. On the other hand, classification of Fusarium species has still a number of open question marks than need to be solved and should require the use of all species recognition methods in an integrated approach. The G. fujikuroi species complex. Gibberella fujikuroi (Sawada) Ito in Ito e K. Kimura has been considered for a long period the teleomorph of several Fusarium species, morphologically placed by Nelson et al. in the Liseola Section.4

Molecular Detection of Foodborne Pathogens

Within this complex, Nelson et al.4 comprised four anamorphic species, including the maize pathogens, F. moniliforme, F. proliferatum, and F. subglutinans, and a minor species, F. anthophilum. On the other hand, Gerlach, and Nirenberg3 identified ten species in Section Liseola and adopted the name of F. verticillioides (Sacc.) Nirenberg (teleomorph Gibberella moniliformis), instead of F. moniliforme, as generally accepted by the research community of Fusarium.13 As for many other fungal phyla, taxonomic results based on MSR, BSR, and PSR (see Section 40.1) have started recently to be compared also for G. fujikuroi species complex. For what BSR is concerned, many reports have now clarified that G. fujikuroi species complex includes at least 11 different biological species or mating populations (MPs): MP-A (G. moniliformis, anamorph F. verticillioides), MP-B (G. sacchari anamorph F. sacchari), MP-C (G. fujikuroi anamorph F. fujikuroi), MP-D (G. intermedia anamorph F. proliferatum), MP-E (G. subglutinans anamorph F. subglutinans), MP-F (G. thapsina anamorph F. thapsinum); MP-G (G. nygamai anamorph F. nygamai); MP-H (G. circinata anamorph F. circinatum), MP-I (G. konza anamorph F. konzum), MP-J (G. gaditjirrii anamorph F. gadtijirrii) and MP-K (G. xylarioides anamorph F. xylarioides).2 The results of sexual crosses, integrated with morphological observations and molecular data by using Random amplification of polymorphic DNA (RAPD), Amplification fragment length polymorphism (AFLP), Restriction fragment length polymorphism (RFLP), and DNA sequencing,6,14–17 have shown that the results of the three classification methods (biological, morphological, phylogenetic) are largely congruent. However, phylogenetic analyses carried out by O’Donnell et al.14 using several genes among which β-tubulin and calmodulin revealed 46 species in the G. fujikuroi complex, 23 of them new to science. Among the 46 species, the 11 species identified by using biological species concept have been reported identical to the phylogenetic species which indicates that phylogenetic approach can provide the same information as biological approach and that other MPs need to be still identified. The F. graminearum species complex. The recent re-classification of F. graminearum (teleomorph, G. zeae), a worldwide pathogen of wheat and maize, is controversial. F. graminearum produces several mycotoxins, mainly trichothecenes, which are tricyclic sesquiterpenes that have been strongly associated with chronic and fatal toxicoses of humans and animals, and zearalenones, which have estrogenic activity.7 F. graminearum produces multiple trichothecene analogues, in particular deoxynivalenol (DON) and nivalenol (NIV) and their acetylated derivatives, 3-acetil-DON (3A-DON) and 15-acetilDON (15A-DON). Within this species, strains differ in their trichothecene production profiles; some strain produce DON, some produce NIV, and others produce DON and NIV. Such chemotype diversity within F. graminearum is a result of loss of gene function. DNA sequence-based phylogenetic analysis of F. graminearum field isolates from six continents delineated eight phylogenetically distinct lineages that were considered biogeographically structured.18 Among the lineages, lineage 7 was considered the most geographically widespread lineage, predominating on wheat and maize in North and South

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Fusarium

America, and in Europe and producing primarily DON.18 The lineages were considered genetically isolated because each was reciprocally monophyletic within genealogies when the six nuclear genes were analyzed individually and when all six were analyzed together. Further studies based on DNA sequence polymorphisms from 11 nuclear genes and three intergenic regions led O’Donnell et al.19 to describe nine lineages within F. graminearum clade and to elevate these lineages to the rank of species. Finally, Starkey et al.20 described two novel species within the F. graminearum species complex based on phylogenetic analyses of multi-locus DNA sequence data of 13 genes. However, not all Fusarium researchers agree with the division of F. graminearum into multiple species. Some authorities considered the lineages to be subspecies rather than species.2 This opinion was supported by the finding that in general, isolates from partially inter-fertile phylogenetic species tend to have AFLP band identities in the range of 40–65%, and that F. asiaticum and F. graminearum have AFLP band identities of 50%. Moreover, only three of the nine lineages show conidial morphology traits useful for differentiating between them, and there is no correlation between lineages and specific mycotoxin profile.18,19,21 Finally, sequencing a portion of tri101 gene of 400 strains of the F. graminearum species complex, Leslie et al.22 generated both a phylogenetic tree and a genetic network that led to clearly state by the authors that “there is only a single species within F. graminearum/G. zeae.” Due to these apparently contradictory data, additional studies are necessary to determine whether the different lineages of F. graminearum represent distinct phylogenetic species or subspecies lineages. Such studies should also provide insight into the practical implications dividing F. graminearum into multiple species with respect to disease management, quarantine regulations and plant breeding strategies and to better understand the ecology, epidemiology, and population dynamics of F. graminearum species complex. The third case, Fusarium oxysporum: a species complex? F. oxysporum is a plant pathogen causing a wide range of plant diseases mainly related to vascular wilts. However, within the species, many populations isolated mainly from soil have been shown as nonpathogenic and they are used as bio-control agents against several diseases also caused by Fusarium species.23 Morphologically, these strains cannot be differentiated from pathogenic strains, although a wide genetic diversity of the population originated from soils has been reported.23 On the other hand, the majority of the isolates causing vascular wilts are specific for a certain host plant. From taxonomic point of view, these strains have differentiated from each other on the basis of pathogenicity as formae speciales. Therefore, the identification of these strains traditionally involves tests of pathogenicity with the appropriate hosts, which are time consuming and can require several months for some formae speciales. Moreover, since pathogenicity is not an ancestral character, taxonomic distinctions of strains based only on this are not reliable from an evolutionary point of view and formae speciales should not be considered monophyletic in origin. On the other hand,

the basis for formae speciales names need not be grounded in traits that are monophyletic in origin24,25 in order to avoid mistakes in breeding for resistance, and to set up inappropriate quarantine measures.2

41.1.2  Biology and Pathogenesis As stated above, several Fusarium species can produce a diverse range of toxic metabolites which may contaminate foods and animal feeds of plant origin,7 but only a small number are considered important for human and animal health. Among these are trichothecenes and fumonisins, molecules very resistant to the transformation processes.26 Trichothecenes. The trichothecenes are a family of epoxysesquiterpene metabolites. Based on their chemical structures, the trichothecenes produced by Fusarium species are conveniently sub-divided in two classes: type A and type B, of which T-2 toxin and DON are respectively the most common mycotoxins. The epoxide ring in these molecules is remarkably stable and it is an essential feature for the expression of toxicity. Type A and type B trichothecenes are distinguished by the absence or the presence of a carbonyl group at the C8 position.7 Trichothecenes are known to act as translational inhibitors in eukaryotic ribosomes,27 but these compounds can be also virulence factors in the infection of wheat plants by their producing fungi.7 Among the fusariotoxins, the T-2 toxin in one of most acutely poisonous, produced mainly by F. sporotrichoides and F. langsethiae, two pathogens of cereals commonly occurring in environment characterized by cold climate. At a lesser extent, F. armeniacum, F. sambucinum sensu strictu and F. equiseti have been also reported as T-2 producers. The illness associated with the consumption of cereals highly contaminated by T-2 includes nausea, vomiting, necrotic lesions in the mouth and throat (making it difficult to eat), severe hemorrhagic diarrhea and hemorrhage in many of the body organs. T-2 toxin is also a potent immunosuppressant because it causes irreversible damage to the bone marrow, leading to a characteristic reduction in white blood cells (aleukia). Among type B trichothecenes, DON is much more common than the T-2 toxin in cereals such as wheat, barley, oat, maize, and at a lesser extent, rice. It is produced primarily by F. graminearum and F. culmorum, the causal agents of red ear rot in maize and head blight in wheat. DON is also known as vomitoxin because of its potent emetic properties and its action as a feed refusal factor. Although much less poisonous as an acute toxin than T-2 toxin, DON may still have a biological effect at very low concentrations. There have been instances in Asia of illness in humans, such as vomiting, nausea, dizziness, and headaches, associated with the consumption of cereals contaminated with DON and possibly much lower concentrations of other trichothecenes. There are considerable annual variations in DON contamination, which usually shows a direct correlation with the incidence of head blight disease in the field caused mainly by F. graminearum. Differently than the T-2 toxin, DON was shown to inhibit translation in Arabidopsis cells without the induction of a defence response.28 The acute phytotoxicity

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of trichothecenes and their occurrence in plant tissues also suggest that these mycotoxins play a role in the pathogenesis of Fusarium on plants. Several groups have investigated the role of trichothecenes in a number of plant diseases by generating trichothecene nonproducing mutants. For example, DON biosynthesis was blocked by disruption of tri5 in G. zeae, which is pathogen of barley, maize, rice, wheat, and other grains. The virulence of two trichothecene nonproducing mutants was significantly reduced in tests in the growth chamber of wheat seedling blight and head scab.29 Trichothecene nonproducing mutants were less virulent than the trichothecene producing parent in their ability to cause head scab. This evidence confirms that trichothecene toxins are probably virulence factors in Fusarium head blight of wheat. Fumonisins. Fumonisins are a family of mycotoxins produced mainly by F. verticillioides and F. proliferatum, which are two very important pathogens of a wide range of agriculturally important plants worldwide. Several other species, almost all included in the G. fujikuroi species complex, and a single strain of F. oxysporum has been reported to produce fumonisins.7 However, for the number of strains reported for each of these species, the fluctuant levels of their production, and the limited occurrence that these species have on plants of agro-food importance, the risk for human and animal health due to fumonisins are related mostly to F. verticillioides and F. proliferatum. Fumonisins can cause the death of horses by massive necrosis of brain tissue (equine encephalomalacia, ELEM), that of pigs by chronic accumulation of fluid in the lungs (porcine pulmonary edema, PPE), and that of rats by necrosis of the liver.7 Moreover they may also be the etiological agents of esophageal carcinoma (EC) in humans30 and in 1993 the International Agency for Research on Cancer (IARC) evaluated these toxins as Group 2B carcinogens.31 Finally, fumonisin contamination of maize has been associated to high incidence rates of neural tube defects in Guatemala, Transkei, and United States, due to the fumonisin capability to cause folate deficiency.7 Fumonisins possess a sphingoid-like long chain base structure and are potent inhibitors of the sphinganine Nacyltransferase, an ER-localized enzyme.32 Fumonisin induced disruption of sphingolipid metabolism is an important part in the cascade of events leading to altered cell growth, differentiation, and cell injury observed in vitro and in vivo. In spite of the demonstrated phytotoxicity of fumonisins in several publications,33 their role in pathogenesis and other aspects of Fusarium ecology is unclear. An association between the production of fumonisins and high levels of virulence on maize seedlings has been detected.34 In more recent reports, a significant positive correlation between leaf lesion development on maize seedlings and the production of FB1 is shown by using wild-type isolates of F. verticillioides.35 Moreover, an inverse correlation between root weight/stalk height and the amount of FB1 in seedling roots is demonstrated.35 Therefore in these works, fumonisin nonproducing strains do not cause leaf lesions and/or have significantly less effects on root weight and stalk height. These works suggest a role as virulence factors for fumonisins. Another report supports the hypothesis for which

Molecular Detection of Foodborne Pathogens

the production of fumonisins is not required for F. verticillioides to infect maize and to cause ear and stalk rot. In particular in this work fumonisin nonproducing mutants obtained by disrupting fum1, a key gene for fumonisin production,7 were used to inoculate maize plants in the field36 and were able to colonize their hosts as successfully as the parental wildtype strain. However these results are made inconclusive by the fact that fumonisin levels were almost equally high in maize artificially inoculated by the nontoxigenic strain and by the toxigenic control, unless the percentage of the nontoxigenic strain exceeded 75–90% of total inoculum. This suggests that the endophytic population of toxigenic Fusarium strains, not eliminable under field conditions, is induced to produce more fumonisins in conditions of high competition,36 and does not allow to evaluate the eventual phytopathogenic performances of the nonproducing strain and in the end the role of fumonisins as virulence factors. Also the F. verticillioides strains isolated in Nepal carry a point mutation on fum1 coding sequence and do not produce fumonisins.37 They are able to cause maize stalk rot under greenhouse conditions38 and maize ear rot and ear infection in small scale field tests.37 These results seem to indicate that fumonisins might be negligible as virulence factors under standard environmental conditions at least in the interactions tested (i.e., for specific maize hybrid-fungal isolate pairs). Fumonisin nonproducing strains associated with banana fruits produced in Central and South America39,40 have been characterized molecularly in a recent work.41 The FUM cluster is absent in these Fusarium isolates; complementation allows fumonisin production and confers pathogenicity on maize seedlings.41 This result indicates that fumonisin production by F. verticillioides may be required for development of foliar disease symptoms, at least on some very susceptible maize genotypes. However, these isolates seem to belong to a genetically drifting sub-­population of F. verticillioides or even to a new, still undescribed species though biologically, morphologically, and phylogenetically close to F. verticillioides,40 thus limiting the possibility to apply these results to the whole F. verticillioides species. A wide number of further fusariotoxins has been shown to have dangerous effects in both animal and plant systems. Among these, zearalenons, nonsteroidal estrogenic metabolites, the cyclic depsipeptides beauvericins and enniatins, the gibberellins, fusaric acid and moniliformin, are the most common. Readers are referred to the excellent book of Desjardins,7 for a complete review on the Fusarium mycotoxins. In this chapter, we underline the importance of only trichothecenes and fumonisins, since the occurrence of these mycotoxins is under strict control by the legislation worldwide due to their impact on human and animal health, their genetic pathways have been well elucidated, and a number of diagnostic tools detecting the genes involved in trichothecene and fumonisin production have been developed. Given the wide range of the toxigenic Fusarium species contaminating maize and wheat, and the diversity of the mycotoxins they can produce,42–46 an assessment of risks posed by maize and wheat consumption by humans and animals has to be determined by correct identification

Fusarium

of Fusarium species with modern, fast, and easy molecular tools. Moreover, a challenge for researcher of the field will be to investigate interactions between the different metabolites that can be accumulated on maize and wheat. The toxic effects that result from combinations of two or more mycotoxins should provide insight into potential consequences of such interactions on human and animal health.

41.1.3  Diagnosis of Fusarium Detection and control of Fusarium species in planta is crucial to prevent mycotoxins from entering the human food and animal feed. Since different Fusarium species have different mycotoxin profiles, an accurate determination of the Fusarium species present is critical to predict potential toxicological risk. The conventional approach to differentiate and detect fungal species in infected plants involves isolation and growth of the fungus in axenic culture followed by identification based upon morphological characteristics,3,4 such as shape, size, and presence or absence of macroconidia, and colony morphology. However, this approach does not provide early detection, can require considerable expertise to distinguish between closely related species,2 can be time consuming, due to long incubation periods, and can be applied only to viable mycelium. In order to overcome these disadvantages, nucleic acidbased methods have been developed to rapidly detect and identify mycotoxigenic fungi. The use of polymerase chain reaction (PCR)47 for fungi detection has replaced cumbersome and time-consuming microbiological analysis, by amplification of specific genomic markers rather than growing the living organism under study. This approach may be faster than culture-dependent methods, and could especially be of value for the rapid detection of slow growing toxin-producing species in food samples. Molecular methods detect sequence differences in DNA or RNA molecules that exist between organisms. Presence of specific PCR product can be detected by gel electrophoresis, hybridization, and sequencing, the main methods to detect nucleic acid differences. Phylogenetic analyses, therefore, become relevant to identify specific nucleic acid regions that can be used for distinguish between species. Phylogenetic analyses of Fusarium,6,48,49 a number of potentially diagnostic sequence motifs were retrieved. PCR-based diagnostics using species-specific oligonu­ cleotides are used in most molecular detection protocols designed to detect single species. In conventional PCR-based assays, specificity can be incorporated in one or both primers, which yield a single band when the target species is present. Specific primers can amplify fragment considered diagnostic that are not part of known coding regions or are part of gene with unknown functions. Anonymous fragments are DNA sequences of unknown function which are used as specific markers for a species or a group of interest, as a unique binding molecule must be detected in a randomly generated set.

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In the early 1990s, random amplification of polymorphic DNA (RAPD) was one of the most common methods for developing genetic species- and population-specific markers for fungi. Today, RAPDs have been replaced by a more robust method, amplification fragment length polymorphism (AFLP). The technique known as sequence characterized amplified region (SCAR) allows sequencing of unique anonymous fragments to design new PCR primers that may or may not maintain the desired specificity. The majority of DNA sequence data used for taxonomy of fungi comes from the nuclear ribosomal operon-like and the regions comprising the ribosomal operon internal transcribed spacers (ITS). The DNA coding for the 18S and 5.8S RNA subunits of eukaryotes are generally too conserved to be used for species delineation or diagnostics. Similarly, the ITS of ribosomal genes do not always provide species level resolution. Despite this, the high number of copies of ITS per cell makes it an attractive target for diagnostics and allows it to be detected with great sensitivity. Species specific assays could be designed also based on ubiquitous, single-copy gene regions such as elongation factor 1-α (TEF-1 α), calmodulin (CaM), beta-tubulin (Ben-A). An alternative strategy that deals specifically with mycotoxigenic fungi is to design primers complementary to mycotoxin biosynthetic genes.50 This approach has the advantage of detecting the fungi that potentially produce the toxins, but it does not take to account that phylogeny of toxin biosynthetic genes is sometimes different from the phylogeny of the species51 and that nonfunctional toxin genes can be present in fungi that do not produce the toxins. As a consequence, numerous assays based on PCR have been developed for identification and differentiation of Fusarium. Most assays are based on ­anonymous frag­ ments,52–54 rDNA, including ITS,52,55 or structural genes56–59 including genes responsible for mycotoxin production.58,60–62 Other molecular methods include RAPD analysis,63 AFLP,64 RFLP,65 specific diagnostic PCR primers,52,62,66–69 SCAR polymorphisms,70,71 DNA sequencing,14,72 the use of secondary metabolites as a taxonomic tool73,74 and the recent use of quantitative real-time PCR assays75–77 have greatly enhanced the ability to detect and distinguish between Fusarium species. The most common secondary metabolites produced by Fusarium species are enniatins, fumonisins, fusaric acid, moniliformin, and trichothecenes.7 The most economically significant are fumonisins and trichothecenes. Trichothecenes occur primarily in small grain cereals, such as wheat, barley, oats, and rye, whereas fumonisins occur mainly in maize. Fusarium graminearum, F. culmorum, F. sporotrichioides, F. poae and F. equiseti are the most important trichothecene-producing species in cereals; F. verticillioides and F. proliferatum are the most important fumonisin-producing species. The toxicity and widespread occurrence of trichothecenes and fumonisins have lead to significant efforts over the past decade and a half to understand the genetics and biochemistry of their biosynthesis. Many of the genes and enzymes

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involved in trichothecene and fumonisin biosynthesis have been identified in significant detail. Genes that encode enzymes required for formation of a mycotoxin or family of structurally related mycotoxins are typically located adjacent to each other in a gene cluster. In general, mycotoxin biosynthetic genes are not expressed constitutively.78,79 Their expression can be detected before mycotoxin production is detected by chemical analytical methods.80,81 For this reason, expression of these genes can be used as an early indication of mycotoxin biosynthesis. However, only some genes of a given mycotoxin biosynthetic pathway can be regarded as key genes, whose activation is directly coupled to mycotoxin biosynthesis. Detection of multiple species that produce the same mycotoxin, rather than a specific species, has also been attempted using primers developed from sequences of toxin cluster genes. With the growing genetic information that is now available, microarray techniques have gained a great deal of attention because of their potential to rapidly and effectively discriminate and identify sequences of different genes in various species, bypassing the need for time-consuming plating and potential misidentification due to lack of diagnostic morphological characters. Microarray technology is a rapid and cost-effective method that has potential for parallel analysis of a high number of target organisms, such as Fusarium species from food. Nicolaisen et al.82 made an oligonucleotide array by with the intention to detect and differentiate nine of the most important Fusarium species occurring on cereal grain, exploiting differences in ITS sequence. Kristensen et al.83 described a DNA microarray for easy and fast detection and identification of 14 trichothecene- and moniliformin-producing Fusarium species. Schmidt-Heydt and Geisen84 developed a microarray which covers most of the known relevant mycotoxin biosynthesis genes. The microarray carries oligonucleotides probes for of the fumonisins, aflatoxins, ochratoxins, trichothecenes (type A and B) and patulin biosynthetic genes. For trichothecene-producing Fusarium the core biosynthetic gene cluster in producing F. sporotrichioides (type A) and F. graminearum (type B) have been spotted. 41.1.3.1  Fumonisins-Producing Species Fumonisins are a group of mycotoxins produced by species within the Gibberella fujikuroi complex species and some strains of fungi within the F. oxysporum complex. Fumonisins belong to a group of fungal metabolites that are structural analogues of the sphingolopid metabolites, sphingosine, and sphinganine,85 able to disrupt sphingolipid metabolism by inhibiting the enzyme ceramide synthase.86 At least 28 fumonisin analogues are known, but the most abundant natural forms are fumonisins B1, B2, and B3.87 In particular, fumonisin B1, containing a keto group at the C-8 position of the trichothecene ring, has been associated with neural tube defects in experimental rodents and may therefore be involved in cases of spina bifida in some humans populations.88 The toxin has been assigned a 2B carcinogen (probably carcinogenic to humans) status by the IARC.31

Molecular Detection of Foodborne Pathogens

Production of fumonisins is restricted to a limited number of Fusarium spp.89 The species reported to produce the highest levels of FBs are F. verticillioides, F. proliferatum, and F. nygamai.90 Among these, F. verticillioides is generally recognized as the most important ear and stock rot pathogens of maize worldwide and responsible for frequent fumonisin contamination.33 Consumption of fumonisin-contaminated feed can cause several fatal animal diseases, including cancer in rodents,91 and there are epidemiological associations between the consumption of F. verticillioides-rotted maize and human esophageal cancer in some regions of the world where maize is a dietary staple.30 Genes involved in the biosynthesis and regulation of fumonisins are organized in a cluster within the genome, as is the case for biosynthetic genes for many other secondary metabolites produced by filamentous ascomycetes: trichothecenes,92 and gibberellins93 in Fusarium, aflatoxin/ sterigmatocystin,94 and lovastatin95 in Aspergillus. To date, fumonisins biosynthetic genes have been mapped to a 42.5-Kb locus in the F. verticillioides chromosome 196 which is absent in F. graminearum, but the region flanking the cluster map to F. graminearum genomic contig 1.159.97 Some species of Fusarium (e.g., F. dlamini, F. napiforme, F. subglutinans and F. thapsinum) in which FUM genes were not detected were reported to produce fumonisins in previous studies,98–100 which could be explained by a difference in ability to produce fumonisins among strains within a species or by misidentification of species. The polyketide synthase gene (fum1) was the first fumonisin biosynthetic gene to be cloned and is the anchor of a cluster of 16 co-regulated fumonisin biosynthetic genes:96,101 12 genes encode biosynthetic enzymes, two encode transporters, and two encode proteins putatively involved in self-protection. A majority of the cluster genes have been functionally characterized and have been found to play a role in fumonisin biosynthesis.102–104 The roles of some of the genes in the cluster have been confirmed by gene deletion analysis96,103–106 and heterologous expression.107 The cluster also encodes an oxoamine synthase (fum8), a C-3 carbonyl reductase (fum13), and cytochrome P450 monooxygenases and other enzymes that catalyze oxygenations at C-5 (fum3), C-10 (fum2) and at an undetermined site (fum6).108 Four genes (fum7, fum10, fum11, fum14) are required for tricarballylic acid esterification. At the opposite end of the cluster from fum1 are genes encoding a transporter protein ( fum19) and for two proteins (fum17 and fum18) with predicted functions in fumonisin self-protection and sphingolipid metabolism. Deletion of either fum1, fum6, or fum8 blocked accumulation of all fumonisins, indicating that these genes are required for fumonisin production.105,106 Finally, Brown et al.101 identified a previously undescribed gene, fum21, located adjacent to the fumonisin polyketide synthase gene, fum1. This gene is a regulatory gene and it is predicted to encode a Cys6type transcription factor. The fum21 deletion mutants do not produce fumonisins and do not express the other genes in the FUM cluster.101

Fusarium

The fumonisin gene cluster was sequenced and annotated also in F. proliferatum by Waalwijk et al.109 which found 19 genes involved in biosynthesis and regulation of the toxin, demonstrating full synteny between F. proliferatum and F. verticillioides, although the flanking regions were completely different. Finally, the FUM cluster has also been identified in an unusual strain of F. oxysporum.110 The F. oxysporum cluster is very similar to the F. verticillioides cluster, but the region flanking the cluster are different.110 Mycotoxin biosynthetic genes can be useful targets to assess the presence of mycotoxigenic fungi in grain and grain-based food and feed. In developing molecular detection methods, genes directly involved in fumonisin biosynthesis are good targets to assess the presence of fumonisin-producing fungi and their ability to produce the toxin in foodstuffs. Recently, PCRbased assays were described that used FUM gene sequences. Bluhm et al.58,60 developed a multiplex PCR assay for the simultaneous detection of fumonisin (based on polyketide synthase, fum1 gene) and trichothecene-(based on a transcription factor, tri6 gene) producing Fusarium sp. in cornmeal. This qualitative tool was upgraded to a semi-quantitative method60 that permitted a correlation between the amount of mycotoxin biosynthetic genes detected by real-time PCR and the capacity to quantify the level of the fungi responsible of toxin production from naturally contaminated field samples of barley and maize. Similarly, Sanchez-Rangel et al.111 developed a fum1-based PCR method that relied on the intensity of PCR fragments on agarose gels. Although this method is not quantitative, it showed a good correlation between fumonisin content and intensity on gel in both inoculated and natural maize samples. For the selective identification of toxigenic isolates of F. verticillioides, Gonzalez-Jaen et al.112 demonstrated that genes fum1 (=fum5), fum6, and fum8 were only present in F. verticillioides, F. proliferatum, F. fujikuroi and F. nygamai, and derived primers from the sequence representing the β-ketoacyl reductase domain within the fum1 gene. Sanchez-Rangelet al.111 reported similar results with a different pair of primers for the same part of the fum1 gene. In contrast to Gonzalez-Jaen et al.112 correlation of PCR results with in vitro fumonisin production of F. verticillioides revealed a number of cases in which a fum1 gene was detected but toxin concentrations produced were negligible or very low. Another group specific assay for fumonisin producers was published by Bluhm et al.58 The system was based on the fum1 gene sequences of F. proliferatum and F. verticillioides and was applied to the detection of these fungi in artificially contaminated cornmeal in a multiplex PCR assay. Potential producers of fumonisins were detected together with potential trichothecene producers (tri6 gene). Recently, a quantitative detection tool, a quantitative PCR (TaqMan), was developed by Waalwijk et al.113 to enable the monitoring of fumonisin-producing fungi (F. verticillioides, F. proliferatum, F. nygamai and F. globosum) in food and feed commodities targeting gene fum1. On the other hand, because the mycotoxin production profile of some fungal species have been clearly demonstrated,

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the possibility to determine the species of an unknown fungal isolate based on its production profile coincides with the identification of toxicological risk. Searching a FUM gene to assess presence of fumonisins producers has the same meaning of searching their species-specific DNA traits. Attempts have been made to use PCR to study the mycotoxigenic potential of fumonisin producing species because it is well established that only some isolates can produce the toxin in vitro and in planta. Based on sequence variation within the intergenic region (IGS) of the rRNA gene between fumonisin producing and non producing isolates of F. verticillioides, Patiño et al.114 set up a highly sensitive PCR system for toxigenic isolates from different geographical regions and hosts. In 1998 Grimm and Geisen compared nucleotide sequences of the ITS1 region within the rRNA genes of typical fumonisin-producing Fusarium species with those of species that do not typically produce that toxin. A pair of PCR primers and a biotinylated probe were designed to identify fumonisin producers (F. verticillioides, F. proliferatum, F. nygamai and F. napiforme) by a microplate-based enzyme linked oligosorbent assay (ELOSA). Later Patiño et al.114 developed a highly sensitive PCR assay for identification of toxigenic isolates from different geographical regions and hosts, based on sequence variation within the IGS region of the rRNA gene between fumonisin producing and nonproducing isolates of F. verticillioides. When closely related species are able to produce more than one toxin and they possess specific mycotoxin profile, it is important to accurately determine their identity in order to have more precise information about the toxicological risk. For this purpose, Mulé et al.59 designed specific primer pairs to identify F. proliferatum (fumonisin, beauvericin, fusaproliferin, and moniliformin producer), F. subglutinans (beauvericin, fusaproliferin, and moniliformin producer), and F. verticillioides (fumonisin producer), by comparing the calmodulin gene sequence from the three species. Previously, Möller et al.115 used RAPD based markers as sequence source to develop PCR-based detection systems for F. verticillioides and F. subglutinans, which were analyzed in maize in multiplex PCR where primers for detection of both species were combined. Another approach using genomic markers of unknown function was applied by Murillo et al.116 who used the sequence of shotgun fragments of genomic F. verticillioides DNA to set up a primer pair which enabled amplification of a 1600-bp PCR product with DNA of that species. However, in a later study Möller et al.115 tested the primers developed by Murillo et al.116 with DNA from other species from the G. fujikuroi complex and found cross reactivity with isolates of mating populations B, C, D, E, F, and also with F. nygamai and F. oxysporum. 41.1.3.2 Trichothecenes Producing Species The trichothecenes constitute a family of more than 60 sesquiterpenoid metabolites produced by several genera of fungi. The term trichothecene is derived from trichothecin, which was the one of the first members of the family identified. All

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trichothecenes contain a common 12,13-epoxytrichothene skeleton and an olefinic bond with various side chain substitutions. To date more than 200 trichotechenes have been described,117 and they can be divided into four groups according to their chemical stuctrure: type A trichothecenes have a hydrogen or ester side chain at the carbon atom 8 (C-8) (T-2 toxin, HT-2 toxin, 4,15-diacetoxyscirpenol, neosolaniol), type B trichothecenes have a ketone at C-8 (DON, NIV, and their acetylated derivatives), type C trichothecenes have an additional 7,8-epoxide and are a minor group of molecules not produced by Fusarium; type D trichothecenes have a structurally diverse macrocyclic ring between C-4 and C-15 but are also not produced by Fusarium.118,119 Their occurrence as contaminants in crops and therefore their impact on human health is responsible for the interest in these mycotoxins. Type A and type B trichothecenes exhibit acute toxicity, causing vomiting and feed refusal; whereas type B trichothecenes are implicated in more chronic toxicoses, resulting in extensive hemorrhage, a general inflammatory response and hematological toxicities. They also show inhibitory effects on eukaryotic protein biosynthesis and mitochondrial function in vitro120,121 and in vivo122 and immunosuppressing effects at low concentrations.123 Trichothecenes are produced by Acremonium (Cephalos­ porium), Cylindrocarpon, Dendrodochium, Myrothecium, Trichoderma, and Trichothecium, but Fusarium is the genus of most concern because it can produce trichothecenes in edible crops. Trichothecene producing species of major concern are F. graminearum, F. culmorum, F. cerealis, F. poae and F. equiseti.7 These species are divided into two main clades, one producing type A and the other producing type B trichothecenes (see Section 41.1.2). Different chemotypes have been identified based on type B trichotechene production. Among strains of F. graminearum and F. culmorum, three chemotype have been described. Strains with the NIV chemoptype produce NIV and 4-acetylNIV (4A-NIV); strains with the 3A-DON chemotype produce DON and 3A-DON; and strains with 15A-DON produce DON and 15A-DON. Moreover, some Fusarium isolates have been reported to produce both DON and NIV and were described as unknown chemotypes.51 Distinguish between different chemotypes is important because differences in the pattern of hydroxylation and acetylation of trichothecenes can result in marked differences in toxicity and biological activity.124 On the other hand, the taxonomic re-evaluation of F. graminearum in the last decade (see Section 41.1.2), the most important type B trichothecene producer, increase the importance to clarify relationship between chemotypes and species of F. graminearum species complex at both chemical and molecular levels. As the most effective control strategy for mycotoxins is prevention of fungal infection and toxin production in the field and in storage, the understanding of molecular biology of trichothecene production should help the development of practical and specific controls. In recent years, the rapid advances in the molecular genetics of filamentous fungi have opened the way for detailed genetic analysis of trichothecene biosynthesis in Fusarium species.

Molecular Detection of Foodborne Pathogens

It would be optimal to develop a reliable PCR assay that could distinguish between F. graminearum strains with DON and NIV chemotypes and relate this assay to the mycotoxins quantification, as suggested by Li et al.125 To this respect, Jennings et al.46 developed a PCR assay based on genes involved in biosynthesis of trichothecenes that could distinguish F. graminearum strains with 3A-DON and 15ADON chemotypes. The trichothecene biosynthetic pathway in Fusarium species begins with a sesquiterpene cyclization catalyzed by the enzyme trichodiene synthase (TRI5), followed by up to eight oxygenations and four esterifications. Hohn and Vanmidlesworth126 were pioneers in molecular genetic studies of trichothecene biosynthesis. They purified trichodiene synthase, the first committed enzyme in the pathway, and isolated the tri5 gene (previously designated TOX5) from F. sporotrichioides.126 In 1993, genomic cosmid clones were successfully used to complement F. sporotrichioides mutants blocked at different stages in the T-2 biosynthetic pathway, suggesting that at least some genes involved in trichothecene biosynthesis (TRI genes) are clustered.127 Subsequent analysis demonstrated the presence of a cluster consisting of 11–12 genes in both F. sprotrichioides and F. graminearum.128,129 Gene-disruption studies have determined that ten of the cluster genes are coregulated and required for trichothecene biosynthesis. Two of the genes encode regulatory proteins, seven encode biosynthetic enzymes and one encodes a transporter gene regulatory genes.97 Three trichothecene biosynthetic genes are located outside the core TRI cluster: the single gene tri101 locus, and the two gene tri1-tri16 were found outside of the tri5 gene cluster.130 The core TRI cluster, the tri1-tri16 cluster, and the tri101 locus each map to different chromosomes.97 The genetics and regulation of trichothecene biosynthesis have been elucidated in detail in F. sporotrichioides127 and F. graminearum.128 Sequencing of parts of the trichothecene gene cluster was done for species in species genetically relates to F. graminearum, such as F. crookwellense, F. culmorum, F. lunulosporum and F. pseudograminearum.97 The sequence of tri5 was used by Niessen and Vogel131 and Schnerr et al.75 to develop a group specific PCR based assay for detection and identification of ­trichothecene-pro­ducing Fusarium species from environmental samples. Several other tri5-based PCR assays have been developed to detect fungi with the potential to produce trichothecenes.132 Quantitative methods based on the tri5, for the quantification of trichothecene producing Fusarium species, have also been described: a competitive quantification system by Edwards et al.133 and a real-time PCR system by Schnerr et al.75 Moreover, Niessen et al.134 developed tri5-based primers for species-specific detection assays for the newly described species F. langsethiae135 and for F. kyushuense, F. poae and F. sporotrichioides. The detection systems made use of the forward primer described in Niessen and Vogel,131 combined with the reverse primers binding specifically to the intron region of tri5 in the respective species. Using the same gene, Strausbaugh et al.136 recently set up a TaqMan™ quantitative

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Fusarium

real-time PCR assay for quantification of F. culmorum in wheat roots and in barley plants. In addition to tri5, other genes from the trichothecene biosynthetic cluster were used to design species- and groupspecific PCR primers. For example, Bluhm et al.58 developed a group-specific assay for genes involved in tri6, which encodes a transcription factor that activates expression of other genes in the cluster. In NIV-producing strains tri13, and tri11 are intact and functional, but in DON-producing strains tri13 and typically tri7 have multiple insertions and deletions that render them nonfunctional. Recent works by Brown et al.92,137 and Lee et al.138 have shown that the genes tri13 and tri7 from the trichothecene biosynthetic cluster are responsible for conversion of DON to NIV (tri13) and acetylation of NIV to 4A-NIV (tri7). Lee et al.138 developed a PCR system for characterization and differentiation of DON and NIV production phenotypes of F. graminearum by designing primers based on sequences differences of tri7 that exist in DON- and NIV-producing strains they are the same for all NIV chemotypes and highly variable for DON chemotypes, due to the presence of a variable number of 11 bp repeated sequence, responsible of tri7 inactivation. An interesting PCR assay was developed by Bakan et al.139 that applied intergenic sequences between the tri5 and tri6 genes to distinguish between high and low DON producing F. culmorum isolates, providing important details on the toxin production capability of F. culmorum strains detected in wheat kernels. A trichothecene-producing specific primer pair hybridising to sequences within the tri13 gene was published by Demeke et al.140 The authors screened samples of wheat, barley, oat sand corn from Canada and found good correlation between PCR detection of F. graminearum biomass and presence of DON in the samples. Also Jennings et al.46 used a primer set directed against tri3, that differentiates whether the DON phenotype was able to produce 3-ADON or 15-ADON. Recently the same approach was used by Quarta et al.141 to set up a multiplex PCR assay, based on tri3, tri5, and tri7 sequences, in order to determine whether wheat seeds were infected with 3ADON, 15A-DON or NIV-producing strains of F. culmorum, F. graminearum and F. cerealis. Various other PCR approaches based on sequence sources of genes not involved in trichothecene biosynthesis pathway were used to detect trichothecene-producing Fusarium species. Schilling et al.52 constructed specific primers on anonymous marker for F. culmorum, F. avenaceum and F. graminearum. Later, the same research group developed primers specific for the maize pathogens F. verticillioides and F. subglutinans.115 Nicholson et al.66 described a method based on SCAR primers for F. culmorum and F. graminearum to detect both pathogens at once. The primers could also be used to quantify the pathogens by competitive PCR. Doohan et al.142 used ­species-specific PCR to analyse the occurrence of F. culmorum, F. poae, F. avenaceum, F. graminearum, M. nivale var. majus and M. nivale var. nivale on different parts of crop plants. Highly specific identification of F. graminearum was demonstrated by Niessen and Vogel143 who used the galactose oxidase gene (gaoA) as a sequence source to design a

species specific PCR assay. Using the same primer pair in a triplex PCR assay, Knoll et al.144 demonstrated the usefulness of the method for detection of trichothecene-producing Fusarium species in contaminated wheat. 41.1.3.3 Surface Plasmon Resonance (SPR) Biosensors Different molecular approaches based on SPR have been recently adopted to obtain detection of toxigenic Fusarium species on cereals. Kai et al.145 showed that DNA-based biosensor are valid and rapid tools for detecting microorganisms. SPR biosensors have been shown to be rapid, label-free, and selective tools to detect PCR products.146,147 These biosensors measure refractive index changes that occur on a metal film providing a signal that is positively correlated with the mass density changes on the metal surface.148 By immobilizing a biological recognition molecule (ligand) on the sensing surface, SPR biosensors provide a tool for real-time biospecific interaction analysis. The choice of a specific single-stranded DNA ligand should allow rapid detection of the target DNA by hybridization with higher selectivity with respect to gelelectrophoresis anlysis. Using this technique, Zezza et al.149 designed a SPR sensor based on DNA hybridization for the detection of F. culmorum on wheat. Further studies in this direction are in progress and seem to be promising in order to apply the same method to further toxigenic Fusarium species colonizing cereals. Another technique related to SPR is SPR imagining (SPRI) technique that has been used to study the hybridization of single-strand DNA of toxigenic fungi belonging to Aspergillus genus isolated from grape.150 SPRI system is proper choice for a real-time monitoring of hybridization dynamics on an array of immobilized oligonuclotide probes because of the high sensitivity in characterization of ultra-thin films adsorbed onto gold or other noble metal surfaces. By this technique, local changes in the reflectivity from a thin metal film describe the hybridization process between the molecules tethered to the surface and those sent in solution in the test chamber. A SPRI-based sensor prototype was used to study DNA–DNA interactions in solution. The biosensor allowed a high number of loci or fungal species to be analyzed at the same time by using suitable gold chips on which different DNA probes were immobilized in array configuration. A 5′ thiol modified DNA probe specific for F. culmorum was immobilized directly onto the gold surface by means of a self-assembled monolayer technique.151 The sensor created proved to be a potential tool for simultaneous analysis of different fungal species, which can be integrated in control programs to prevent toxigenic Fusarium species through-out the food production chain.

41.2  Methods 41.2.1 Molecular Detection of FumonisinsProducing Fusarium spp. in Maize59 Sample preparation. Isolate DNA from ground kernels (100 mg) using “Magbeads Plant DNA Kit” (OmegaBiotek) according to the manufacturer’s instructions. The procedure is based on reversible adsorption of nucleic acids to

586

Molecular Detection of Foodborne Pathogens

paramagnetic beads under appropriate buffer conditions. Disrupt plant tissue and then lyse in a specially formulated buffer containing detergent. Subsequently precipitate proteins, polysaccharides, and cellular debris. Transfer the lysate to a 96-well microplate, adjust binding conditions, and bind genomic DNA to the Mag-Bind® particles. Remove trace contaminants such as residual polysaccharides with one or two rapid wash steps. And finally elute pure DNA in water or low ionic strength buffer. Purified DNA can be directly used in downstream applications without the need for further purification. Detection procedure. The three primer pairs (PRO1/2, SUB1/2 and VER 1/2) listed in Table 41.1, targeting calmodulin gene, amplify PCR products of 585, 631, and 578 bp for F. proliferatum, F. subglutinans and F. verticillioides, respectively. Perform PCR reactions for the three species in separate tubes. Set up the single PCR assay in 25 µl volume: 0.13 µl (5 U/µl) of Taq Gold DNA polymerase (Applied Biosystems), 2.5 µl 10 × Taq Gold polymerase buffer, 7.5 pmol of each outside primer, 12.5 µM of each deoxynucleoside triphosphate, and 0.5 µl (out of 100 µl) of total nucleic acids isolated from food fungal template DNA. Perform PCR amplification as follows: one cycle of 95°C for 5 min; 35 cycles of 95°C for 50 sec, 56°C for 50 sec, and 72°C for 1 min; one cycle of 72°C for 5 min; followed by cooling at 4°C until recovery of the samples. Check presence/absence of PCR products in 1.5% ­Tris-acetate-EDTA-agarose gel and visualize with ethidium bromide and ultraviolet illumination.

41.2.2 Quantitative Detection of FumonisinsProducing Fusarium spp. in Maize113 Sample preparation. Grind freeze-dried material to a very fine powder and sieve to obtain particles of ≤ 1 mM using a cyclotech sample mill (Foss Tecator, Hoganas, Sweden). Isolate DNA from maize samples using the Qiagen DNeasy Plant Mini kit following the protocol, that was earlier applied for wheat samples.77 Suspend 20 mg in 400 µl lysis buffer152 and after 5 sec of sonification these samples are mixed with 200 µl of buffer AP1 of the plant DNeasy kit (Qiagen, Germany). Subsequently, process samples according to the

manufacturers protocol and finally elute DNA with 200 µl of elution buffer AE. Detection procedure. Perform real-time PCR using a MicroAmp Optical 96-well reaction plate and MicroAmp Optical Caps (Applied Biosystems). Use an ABI Prism 7700 Sequence Detection System (Applied Biosystems) to perform the PCR and assess fluorescence. Each amplification reaction consisted of 2 μl of DNA preparations, 1 × real-time PCR buffer (Applied Biosystems), 5 mM MgCl2, 83 nM of the FAM-labeled FUM-probe1, 1.5 U of Hot Gold star DNA polymerase (Eurogentec, Seraing, Belgium) and 333 nM of forward and reverse primer for the target DNA (Taqfum-2F in combination with Vpgen-3R or VertFum-3R (Table 41.2). As an internal control, include in the reaction along with 83 nM of the VIC-labeled PLRV probe, 100 pg of Potato Leaf Roll Virus (PLRV) DNA, forward primer PLRV-F and reverse primer PLRVR (both at 333 nM).77 Thermal cycling conditions consisted of a single cycle of 2 min at 50°C to degrade uracil containing DNA and 10 min at 95°C to inactivate uracil-N-glycosidase, followed by 40 cycles of 95°C for 15 sec and 60°C for 1 min. The assay allows to detect and quantify fumonisin-producing Fusarium spp. (F. verticillioides, F. proliferatum, F. globosum, and F. nygamai) in maize samples.

41.2.3 Molecular Detection of Trichothecenes Fusarium spp. in Wheat141 Sample preparation. Isolate DNA from wheat kernels ground to a powder by a grinder. Start from 100 mg of the sample using the RedExtract-N-Amp Seed PCR Kit (SigmaAldrich Co. St Louis, MO) according to the manufacturer’s instructions. Detection procedure. For the identification of DON producing strains use the primer set 3551H/4056H based on the published tri5 gene sequences. For the identification of 3A-DON or 15A-DON and NIV chemotypes use the primers directed to tri3 and tri7: Tri3F1325/Tri3R1679, Tri3F971/ Tri3R1679, Tri7F340/Tri7R965 (Table 41.3). For verifying the contamination of wheat by 3A-DON, 15A-DON and NIV chemotypes of F. culmorum, F. graminearum and F. cerealis, carry out multiplex PCR experiments

Table 41.1 Identities of Species-Specific Primers Primer

Sequence

SUB1

5′-CTGTCGCTAACCTCTTTATCCA-3′

SUB2

5′-CAGTATGGACGTTGGTATTATATCTAA-3′

PRO1

5′-CTTTCCGCCAAGTTTCTTC-3′

PRO2

5′-TGTCAGTAACTCGACGTTGTTG-3′

VER1

5′-CTTCCTGCGATGTTTCTCC-3′

VER2

5′-AATTGGCCATTGGTATTATATATCTA-3′

Specificity F. subglutinans F. proliferatum F. verticillioides

Cycling Conditions 5′/ 95°C (50′′/ 95°C, 50′′/ 56°C,1/ 72°C) 35× 5′/72°C

Amplicon (bp) 631 585 578

587

Fusarium

using 10–25 ng of fungal DNA in a total volume of 50 µl of 1 × reaction buffer containing 1.5 mM MgCl2, 2 U Taq DNA polymerase (EuroTaq, Euroclone, Lugano, Switzerland), 200 µM dNTPs, 0.2 µM each of the three tri3 primers and 0.1 µM each of the remaining four primers reported above. Perform PCR cycle as follows: one cycle of 94°C for 3 min; 35 cycles of 94°C, for 30 sec, 53°C for 30 sec, and 72°C for 1 min; and 1 cycle of 72°C for 10 min. Check presence/absence of PCR products in 1.5% ­Tris-acetate-EDTA-agarose gel and visualize with ethidium bromide and ultraviolet illumination. The detection limit of the assay on an ethidium bromide-stained agarose gel is about 4 pg of fungal DNA.

41.3 Conclusions and Future Perspectives The increasing worldwide concern regarding food safety led to consider the control of contamination of food and feeds with Fusarium mycotoxins and related producing species a major aim. Therefore, there is a need for developing rapid, sensitive, and inexpensive diagnostic tools for the detection of the main toxigenic Fusarium species, both in the field and after harvest to obtain real-time monitoring data on contamination and better assess food safety. This will result in an enormous cost saving to the farmers as well as to agro-food industry through the prevention and reduction of product recalls and reduce anti-fungal treatment costs. As Table 41.2 Primers and Probes Name

Sequence (5′–3′)

Taqfum-2F Vpgen-3R FUM-probe1

ATGCAAGAGGCGAGGCAA GGCTCTCA/GGAGCTTGGCAT1 FAM-CAATGCCATCTTCTTG

PLRV-F PLRV-R PLRV probe

Internal control AAGAGGCGAAGAAGGCAATCC GCACTGATCCTCAGAAGAATCG VIC-CGAAGACGCAGAAGAGGAGGCAAT

Note: A/G indicates a degenerate nucleotide at this position.

stated in the introduction to this chapter, the main diseases caused by Fusarium species on wheat and maize, two major sources of food worldwide, results from complex species attacks showing a wide array of species biodiversity. As a main consequence, the mycotoxin contamination profile and related toxicological risk can dramatically vary according to the prevailing Fusarium species in the food. Moreover, the strategies for the disease control have to face a range of Fusarium species each with its own ecology, epidemiology, and population dynamics. Therefore, to combine accuracy, reliability, and rapidity in the diagnosis of Fusarium species in cereals is a main challenge for scientists working in the field. The PCR methods largely illustrated in this chapter have shown that a wide number of species-specific DNAbased assays have been designed for several mycotoxigenic Fusarium species, enabling also a direct detection from field samples without the isolation and purification of cultures. Because species concepts for many mycotoxigenic Fusarium species have fluctuated over time, better taxonomic keys and sequence databases will be useful to make identification of pure cultures faster and more accurate. Phylogenetic databases could provide new insights to resolve many of these taxonomic issues and should make it technically feasible to have a single assay that would detect all known mycotoxigenic species. Among the most advanced techniques in applied molecular genetics, the development of the DNA/RNA probe technology is of particular interest. Species differences result in some sequence variations so sequence analysis of some variable and conserved DNA regions (e.g., 28S, ITS-1/ITS2, β-tubulin, calmodulin, and elongation factor) is informative at the intra- and inter-specific level. Since such sequences could be useful to design molecular markers, and as the complete genome sequences of some mycotoxigenic Fusarium species (F. verticillioides and F. graminearum) became available, DNA arrays using these sequences are being developed by public and private laboratories.83, 84 DNA arrays are miniaturized microsystems based on the ability of DNA to spontaneously find and bind its complementary sequence, fixed to solid surfaces, in a highly specific and reversible manner, known as hybridization. They are based on the fact that a single base mismatch between the oligonucleotide on the array and the gene will hybridize

Table 41.3 Primers and Probes Primer Name 3551H 4056H Tri3F971 Tri3R1679 Tri3F1325 Tri3R1679 Tri7F340 Tri7R965

Primer Sequence ACT TTC CCA CCG AGT ATT TT CAA AAA CTG TTG TTC CAC TGC C CAT CAT ACT CGC TCT GCT G TT(AG) TAG TTT GCA TCA TT(AG) TAG GCA TTG GCT AAC ACA TGA TT(AG) TAG TTT GCA TCA TT(AG) TAG ATC GTG TAC AAG GTT TAC G TTC AAG TAA CGT TCG ACA AT

Target Gene

Amplicon Size (bp)

Tri5

525

Chemotype DON

References 141

Tri3

708

15A-DON

153

Tri3

354

3A-DON

153

Tri7

625

NIV

153

588

very weakly compared to a perfect match. DNA arrays are designed to accelerate the hybridization process by labelling the biological sample and hybridizing it to a large collection of immobilized probes, allowing screening for the presence of up to several sequences in a single experiment. They can be very useful to investigate and detect, at single nucleotide level, differences in gene’s sequences of interest. Although hybridization techniques for analyzing DNA have been known to molecular biologists for decades, the keen interest that is now being shown in these new technologies has stemmed from a number of key innovations as the use of a planar solid support, facilitating miniaturization and highdensity analysis, powerful high-resolution detection and the development of high-density spatial synthesis of short DNA probes (oligonucleotides). Recently, a new type of low-complexity oligonucleotide microarray concept, applied to the identification of Aspergillus species has been developed by Bufflier et al.154 This microarray format could be well adapted to detect different toxigenic fungi, including Fusarium toxigenic species, in parallel in food samples. Toxigenic fungi genomic DNA extracted from food can be multiplex PCR amplified to generate biotin-labeled fragments and revealed on biochip OLIgo Sorbent Arrays (OLISATM, Apibio, Francia) microarray by colorimetric detection. Wells are imaged with an automated system and spot density is analysed by calculation of the average grey level of each spot minus that of the average well-bottom background. The associated software allows an automatic analysis of images and provides an easy-to-use final report of results. The presence of multiple toxigenic Fusarium strains could be detected in one step. With increased interest in research dealing with molecular methods, biosensors, automation, and miniaturization, there are high prospects for development of fast, accurate, sensitive, user friendly, and cost-effective methods in food mycology in the near future, including the detection of Fusarium species. The possibility to develop more accurate and fast PCRbased assays on main foodborne Fusarium species is extremely useful for their application in food hazard analyses. False-negative reactions have to be promptly identified by using internal DNA control. Moreover, (i) extensive testing for specificity of target sequences, (ii) optimization of sensitivity in food matrixes by reducing the effects of the inhibiting factors, and (iii) the friendly use for trained operators, are important objects to pursue. Finally, further challenges in detection of Fusarium species include the possibility to combine the great potential of the microarray technology with the current availability of many probes for biosynthetic genes involved in trichothecene and fumonisin pathways. This would allow not only the detection of the main occurring toxigenic Fusarium species directly from field samples by implementing them in DNA biochips, but also to investigate the mycotoxin gene expression. Specific expression profiles of the whole mycotoxins biosynthetic pathways can be generated by RNA microarray allowing the exact prediction and control of food contamination. This kind

Molecular Detection of Foodborne Pathogens

of microarray could be used in the future also for studying the effects of different relevant parameters on the mycotoxin gene expression. In this respect, the use of DNA-based assay for estimating the risk of contamination is severely problematic because (i) the mycotoxins could be present without the fungi, (ii) the fungi can be present without mycotoxins, and (iii) DNA quantification is not ‘‘one gene/one spore’’ since spores of Fusarium species can have multiple nuclei. Thus, it will be important to use an innovative approach for diagnosis of Fusarium species allowing the PCR assay to detect, in field samples, the main occurring toxigenic Fusarium, their relevant mycotoxin genes and determine whether or not such genes are expressed, to reliably assess the real toxicological risks related to Fusarium contamination of food.

References





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42 Penicillium Joëlle Dupont

Natural History Museum of Paris

Contents 42.1 Introduction.................................................................................................................................................................... 593 42.1.1 Classification, Morphology, and Biology.......................................................................................................... 594 42.1.2 Epidemiology and Pathogenesis....................................................................................................................... 595 42.1.2.1 Mycotoxinogenic Foodborne Penicillia........................................................................................... 595 42.1.2.2 Foodborne Penicillia Involved in Human Pathogenesis . ............................................................... 595 42.1.3 Laboratory Diagnosis........................................................................................................................................ 596 42.1.3.1 Conventional Techniques................................................................................................................. 596 42.1.3.2 Molecular Techniques...................................................................................................................... 596 42.2 Methods.......................................................................................................................................................................... 598 42.2.1 Samples Preparation......................................................................................................................................... 598 42.2.1.1 Isolation of Penicillium from the Air............................................................................................... 598 42.2.1.2 Isolation of Penicillium from Food Samples................................................................................... 598 42.2.1.3 Preparation of Fungal Cultures for Toxin Detection....................................................................... 598 42.2.1.4 DNA Extraction from Fungal Cultures for Molecular Identification.............................................. 598 42.2.2 Detection Procedures . ..................................................................................................................................... 599 42.2.2.1 PCR Detection of Penicillium Species............................................................................................ 599 42.2.2.2 PCR Detection of Ochratoxin A (OTA)-Producing Penicillium..................................................... 599 42.2.2.3 PCR Detection of Patulin-Producing Penicillium........................................................................... 600 42.3 Conclusions and Future Perspectives............................................................................................................................. 600 References.................................................................................................................................................................................. 601

42.1  Introduction Penicillium are very diverse and cosmopolite. Their primary habitat is soil or decaying vegetation, but a number of species are clearly associated with food. They are well known as starter cultures for production of cheeses and meat products like salami and dried sausage, but also as a major cause of contamination in food and feeds. In terms of human pathology, unlike bacteria, most of the Penicillium species are hardly able to grow at 37°C and infections are very rare in the human host, with the main exception being P. marneffei, a non-foodborne species. Most of the diseases caused by other Penicillium species, many of which are potentially foodborne, have generally developed after inhalation of spores, not after ingestion of the microorganism. The main risk in ingesting food contaminated by Penicillium is the exposure to important mycotoxins possibly produced by several species. Ochratoxin A (OTA) and patulin are the most significant Penicillium mycotoxins for which regulations are imposed in a number of countries. More often, the risk comes from the ingestion of manufactured products made with contaminated raw materials rather than from food products visibly contaminated,

which are generally rejected by consumers because of their altered organoleptic aspect. The risk is however persistent in countries where no regulations exist, particularly in tropical countries where contaminations are frequently occuring. The difficulty in establishing the incidence of Penicillium species in pathogenesis has been attributed to the lack of rapid and sensitive methods for their identification. The majority of the cases reported are still based on conventional morphological diagnosis resulting in numerous misidentifications. Although complementary growth characters have been proposed1,2 to facilitate the recognition of the species, only highly skilled taxonomists are able to identify Penicillia on phenotypic criteria. The impact of molecular revision of the Penicillium species has just been felt, and presently only the subgenus Penicillium, mostly involved in food and indoor contamination, has been entirely revised in a polyphasic approach combining molecular and phenotypic characters to secondary metabolites profiles,2 providing a solid base for molecular detection. Using molecular approaches, Penicillium morpho species have been shown to enclose two or three phylogenetic species,3,4 the genus Penicillium may contain more than 500 species.

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42.1.1 Classification, Morphology, and Biology Classification. Penicillium are ascomycetous fungi in class Eurotiomycetes, order Eurotiales, family Trichocomaceae. There are at least 250 species of Penicillium presently accepted5 of which only a few species have a complete life cycle, where sexual states are represented either by Talaromyces or Eupenicillium. The remaining species are considered as strictly asexual fungi and may have lost their sexual state independently during evolution.6 The sexual states Talaromyces and Eupenicillium constitute distinct phylogenetic lineages within the Trichocomaceae, in which asexual species are distributed with a clear correlation with their morphology. On the base of the branching pattern of their penicillus, from the simplest to the more complex, asexual species have been classified into four subgenera, Aspergilloides, Biverticillium, Penicillium, and Furcatum.1 The subgenus Biverticillium is phylogenetically related to Talaromyces,6 while the remaining three subgenera are related to Eupenicillium,7,8 showing clear affinities with Aspergillus and related sexual states.9 Morphology. The reproductive asexual apparatus of Penicillium is composed of a brush-like structure called a penicillus that produces exogenous mitotic spores called conidia. The penicillus is composed of a conidiophore bearing phialidic conidiogenous cells directly or on former metulae and branches, determining one or several verticils (Figure 42.1). Monoverticillate penicilli, composed of one whorl of flask-shaped phialides at the top of the conidiophore, characterize the subgenus Aspergilloides. Biverticillate penicilli are typical of the subgenus Biverticillium and terverticillate penicilli of subgenus Penicillium. Some species, having loose penicilli that can not be classified in any of the described categories, are grouped in the subgenus Furcatum. Conidia are produced in dry basipetal chains from the phialides, divergent or in columns visible under

Figure 42.1  Branching patterns of penicilli. (a) Monoverticillate, (b) biverticillate, (c) terverticillate.

Molecular Detection of Foodborne Pathogens

low magnification at the microscope. Conidia are spherical, ellipsoid or short cylindrical, hyaline or greenish, smooth or rough-walled. Slerotia, which are firm and rounded hyphal structures, are produced in some species. The colonies of Penicillia, when grown on suitable media, have generally a velvety, more or less floccose, and sometimes tufted texture. At maturity they are powdery, due to the drywalled conidia, green, greenish blue to blue-grey, surrounded with thin, white to sometimes yellow-orange mycelium. Some species produce typical exudates or pigments, diffusing or not in the agar. On food substrates, the mycelium is often subsurface and only conidia are generally visible, as greenish to blue-grey spots. The sexual spores are produced in globose asci, showing any pore or opening apparatus, contained in fructifications called cleithothecia, cream to yellow-orange colored that are visible to the naked eye. Biology. The genus Penicillium is very diverse in terms of species and range of habitats. Penicillium species are considered to be ubiquitous and opportunistic saprophytes. Most of the described species are primarily soil and decaying vegetation fungi, but quite a number of species are clearly associated with human food supplies. Globally they are nutritionally undemanding, strictly aerobic and able to grow on a wide range of physico-chemical environments, pH, temperature, water availability, CO2 tension, metals, and salts, as well as on complex nutrients. Some species are highly specialized, as fruit pathogens (e.g., P. expansum on apples, P. digitatum and P. italicum on citrus fruits), on low-water content feeds (P. brevicompactum, P. chrysogenum, P. implicatum), and at low oxygen tension (P. roqueforti). Penicillia are mesophilic fungi, with an optimum temperature around 25°C. Most of the foodborne species are psychrotolerant, with the ability to grow at refrigerated temperatures, presenting a risk of contamination for food when refrigerators are not properly managed. On the other hand Penicillium are hardly able to grow at 37°C and are unlikely to be etiologic agents of systemic disease, with the exception of P. marneffei and a few related species (P. piceum, P. rugulosum, P. purpurogenum). Nevertheless, few Talaromyces species that have heat-resistant ascospores may develop on pasteurized products, such as fruit juices, but those species are not known to be toxigenic. Penicillia are fast growing fungi, producing a considerable number of exogenous dry-walled spores that are easily disseminated by air. They are an important component of the bioaerosol. They may settle and colonize all kinds of materials if the relative humidity is available. In food microbiology the water availability of the substrate is expressed in terms of water activity, numerically equal to the equilibrium relative humidity of the air (ERH) expressed as a decimal [aw (food) = ERH (air)/100]. Only few species are able to grow at low aw, i.e., under 0.80. With current understanding of foodborne species,10,11 it is possible to predict which fungi and mycotoxins a given product may contain, when the type of food product and the history of production and storage are known.12

Penicillium

42.1.2 Epidemiology and Pathogenesis 42.1.2.1  Mycotoxinogenic Foodborne Penicillia Unlike Aspergillus and Fusarium, Penicillium are not the prime mycotoxigenic fungi. Most of the Penicillium species are able to produce mycotoxins under laboratory conditions, but they may not do so in food products.13 In some cases, mycotoxins are unstable in food products and P. camemberti and P. roqueforti are unable to produce toxins in industrial conditions.14 OTA and patulin are the most significant of the Penicillium toxins that are submitted for international regulation in food and feeds. The European Food Safety Association (EFSA) has published an updated opinion relating to OTA in food.15 Other important mycotoxins associated with foodborne Penicillia are citrinin, penitrem A, cyclopiazonic, and penicillic acids and the family of the xanthomegnins. Epidemiology. OTA is produced by P. verrucosum and P. nordicum and by some Aspergillus species.2,16 P. verrucosum is mostly found on cereals but also occurs quite frequently on meat products and cheeses. P. nordicum is psychrophilic and was isolated from refrigerated meat products and cheeses. However, cereals are considered as the main source of exposure for humans. It was shown that Aspergillus carbonarius, the main OTA producer in grapes and wines, accumulates high concentrations of OTA in conidia, when the fungus is cultivated on artificial medium. The toxin is rapidly excreted in the medium as soon as conidia germinate.17 OTA is a stable compound that is not destroyed by common food preparation procedures. Temperatures of 250°C are required for several minutes to reduce the toxin concentration.18 Thus raw and processed food commodities can be contaminated by OTA. The maximum concentration recommended in raw cereals and derived products, has been fixed to 5 μg/kg.19 A tolerable weekly intake (TWI) of 120 ng/kg (body weight) has been defined for human exposure.20 Patulin is produced by a number of species in Aspergillus, Byssochlamys, and Penicillium. The most efficient foodborne Penicillium producers are P. expansum on pomaceous fruits and nuts, P. griseofulvum on cereals, P. carneum in beer, wine, rye-bread, and dry meat products, P. paneum on rye-bread and P. sclerotigenum on yams. A regulation has been implemented in some countries for patulin in apple products and the maximum concentration recommended is 50 μg/kg in apple juice.19 The provisional tolerable weekly intake (PTWI) is 0.3 mg/kg body weight (defined by the Joint Expert Committee on Food Additives, JECFA, in its 35th meeting in 1990). Impact on human health. Exposure to OTA has been correlated with Balkan endemic nephropathy and urinary tracts tumors. But convincing epidemiological data are lacking, despite the fact that several publications describe increased OTA blood levels in the population of endemic villages.21,22 The International Agency for Research on Cancer classified OTA as possibly carcinogenic to humans (Group 2B), based on sufficient evidence of carcinogenicity in animals studies and inadequate evidence in humans.23 Patulin is a very

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generally toxic compound, particularly cytotoxic, not classified as carcinogenic in humans (Group 3) by IARC.24 42.1.2.2 Foodborne Penicillia Involved in Human Pathogenesis Clinical cases due to Penicillium have been recently reviewed by de Hoog et al.25 and Lyratzopoulos et al.26 and additional data are available through a computerized literature search for year 2000 onwards. Although Penicillium are phylogenetically related to Aspergillus, they have not the same invasive capabilities, with the exception of P. marneffei, which can cause a fatal systemic mycosis in patients infected with human immunodeficiency virus (HIV). The infection is endemic in tropical Asia.27 Invasive infections due to Penicillium species other than P. marneffei are rare but do exist and currently, P. chrysogenum is particularly suspected to have a significant invasive potential.28,29 Invasive pathologies reported are endocarditis, lung, kidney, and central nervous system infections, endophthalmitis, peritonitis pericarditis, and oesophagitis. It appears that invasive disease can occur in patients who are either immunocompromised or immunocompetent or for which the immune defect was not yet apparent. Penicillium species are responsible for superficial infections including skin infections such as onychomycosis and dermatitis, eye infections such as fungal keratitis and conjunctivitis and otomycosis (refer to De Hoog25 for a description of these pathologies). Allergic diseases are asthma and hypersensitivity pneumonitis. Several occupations in the food production are responsible for hypersensitivity pneumonitis attributable to Penicillium, including cheese and salami workers’ lung.30–32 The majority of the Penicillium species that have been reported from human pathologies are potentially foodborne fungi with the exception of P. marneffei, P. piceum and P. purpurogenum2,10,11 and are ubiquitous and cosmopolite. They are P. brevicompactum, P. chrysogenum, P. citrinum, P. commune, P. decumbens, P. expansum, P. purpurogenum, P. rugulosum and P. spinulosum. Three additional species have caused animals pathologies: P. aurantiogriseum, P. griseofulvum, P. verruculosum. There are very few epidemiological data as most of the cases are isolated. Only few species are able to grow at 37°C (P. chrysogenum, P. purpurogenum and P. rugulosum) and are potentially able to develop invasive infections. P. chrysogenum is a xerophile. It is a very common contaminant occurring on a wide range of food. It is also used as a starter culture for fermented European meat production. P. purpurogenum is not a predominant foodborne fungus, while P. rugulosum has been reported quite often from dried and processed meats. Several species have been isolated from a wide variety of foods, P. citrinum, P. decumbens and P. spinulosum (cereals, nuts, beans, spices, processed meats, fresh vegetables, etc.). Some others are more specialized. P. brevicompactum is xerophile, halotolerant, and psychrotolerant and in response to these physiological traits it is widespread on dried foods

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(grains, cereals, etc.) but also on meat products (hams, salami, etc.) and frequent on refrigerated products (domestic food and stored fruits like apples and grapes). P. commune is primarily found in cheese environments, where it is the principal cause of spoilage, but occurs on various food products and also products with low water content. P. expansum is psychrophile and has a very low requirement for oxygen. It is the principal cause of spoilage of pome fruits and has often been isolated from a wide range of fruits (strawberries, grapes, cherries, peaches, tomatoes, etc.) and nuts.

42.1.3 Laboratory Diagnosis 42.1.3.1  Conventional Techniques There are at least 250 species of Penicillium that have been conventionally diagnosed by morphological characters (macro and micromorphology). The penicillus type alone determines the subgeneric classification of the species. Subgenera are divided into sections according to typical morphological traits. As an example, the texture of the colonies was used to divide the subgenus Biverticillium into two sections, the section Coremigerum grouping species with coremial or synnematous structures, while species with velutinous aspect were classified in section Simplicium. But, because the micromorphology of strains is very similar within each subgenus, other phenotypic characters, e.g., growth at different temperatures or water activities and the presence of pigment and exsudate have been investigated and shown to be of a great interest in Penicillium species delineation.1 The subsequent cultivation method proposed by Pitt1 recommends three different media, malt extract agar (MEA), Czapeck yeast extract agar (CYA) and 25% glycerol nitrate agar (G25N), incubated at 25°C for 7 days, two CYA cultures being also incubated at 5 and 37°C. Morphological observations are done on MEA cultures. Other phenotypic characters are observed on the three media. CYA enhances secondary metabolism, pigments, and exsudates production. G25N which is supplemented with 25% glycerol to reduce the water availability of the medium (0.93 aw corresponding to a relative humidity of 93%, while MEA and CYA are 0.99 aw), is favorable to species able to grow on reduced water activity. Identification keys are proposed for the whole genus1 or restricted to foodborne species.10 Samson and Frisvad2 proposed to use creatine sucrose agar (CREA) in addition to MEA and CYA and to use yeast extract sucrose agar instead of G25N, for identification of species of the subgenus Penicillium. These methods are however difficult to use by nonspecialists, resulting in frequent misidentification that considerably confused the global knowledge of the biology of these microorganisms. To try stopping this chronic problem of misidentification, recommendations have been proposed particularly in the field of mycotoxigenic fungi associated with foods and feeds33 and typical isolates have been defined that can be used as references both for identification and for mycotoxin production.33 In the same way, typical isolates have been defined for each of the 58 species retained in the subgenus Penicillium after a polyphasic taxonomical approach based on phenotypic and

Molecular Detection of Foodborne Pathogens

molecular characters and on secondary metabolites profiles.2 Another reasonable solution would consist of identifying only the subgenus of the incriminated Penicillium strain, as terverticillate Penicillium for example, and focusing on mycotoxin analysis,34 leaving accurate identification to specialist taxonomists of the group. 42.1.3.2  Molecular Techniques Molecular methods based on an accurate phylogenetic reference system offer a big advantage over conventional phenotypic methods for species diagnosis in that they measure stable genotypic characteristics and do not rely on culture and operator interpretation. Molecular evaluation of fungal species originally diagnosed by phenotypic characters, when substantial numbers of strains have been examined, has revealed, in most cases, a proliferation of cryptic species.35,36 An important preliminary work of molecular taxonomy is required in mycology, especially on difficult genera like Penicillium, to define the boundaries of the species prior to suggesting diagnostic tools. The keys to build an accurate phylogenetic reference system are (i) the examination of a sufficient number of strains to evaluate the intra specific diversity (ii) to evaluate accurately the genealogy of the species on a sufficient number of independent genes. It has effectively been shown that single-gene phylogenies do not reflect the underlying organism genealogies because of the stochastic effect of the segregation of variation during speciation and recombination. However the point where different gene genealogies become concordant is a useful place to assign a species boundary.35,37–39 This is the foundation of the Genealogical Concordance Phylogenetic Species Recognition concept (GCPSR)35 that is used in fungal taxonomy. The difficulty of the GCPSR method is to find genes with appropriate evolutionary dynamics as those appropriate genes will vary according to the species investigated. To assess molecular features of the species, both phylogenies of the whole genus establishing species relationships and extensive molecular evaluation of the species to define their genetic variability and their boundaries are necessary. In a large and diverse genus like Penicillium, large phylogenies are essential to decide sequencing strategies for GCPSR assessment of specific questions. This represents a tremendous amount of coordinated work, and there are still difficulties in finding phylogenetically effective genes and enough strains to examine. In the exacting context of searching tools for species diagnosis, the sampling of isolates is decisive. Ex-type isolates have to be considered as taxonomical references, in addition to isolates from diverse habitats to evaluate the intra specific diversity of the species. In that respect, culture collections represent a highly valuable fungal patrimony, but most of the strains have however been diagnosed by phenotypic characters and misidentifications are frequent particularly in difficult species, increasing the sampling difficulty. Isolation of fresh material would be necessary in number of species but is excessively hazardous as their exact habitat is not always understood.

Penicillium

Presently, there is not a complete phylogeny of the genus Penicillium that could be used as a preliminary step toward species diagnosis. Furthermore, from the existing molecular analysis and personal unpublished data, it is obvious that any of the target molecules used until now do not permit the resolution of all the species within the genus because of its important genetic diversity. Thus, molecular diagnostics of Penicillium will turn toward the use of a core molecule allowing a good resolution of most of the species, completed by more specific genes to resolve specific questions. The BarCode of Life initiative aims to sequence short portions of a standardized region of the genome for all forms of life to facilitate rapid low-cost identification by nonspecialists40 (http://www.barcodinglife.org/views/login.php). In this context mycologists have chosen ITS rDNA as the consensus target for barcoding fungi instead of the proposed mitochondrial cytochrome oxydase I (COI). The reasons are the long history of use of ITS rDNA in fungal taxonomy and consequently the numerous sequences already available, the ease of their amplification and their globally good level of resolution, when COI comprises large introns in fungi that make them unapplicable for species diagnosis at a large scale. A number of ITS rDNA sequences of Penicillium are already available in public databases such as GenBank, but the value of the sequence is totally dependant on accurate identification of the organism used. The major problem is that up to 20% of the fungal deposits may be wrongly named.41 The advantage of Barcode of Life Data Systems (BOLD) is that each sequence of reference must derive from a specimen whose taxonomic assignment can be reviewed, ordinarily through linkage to a specimen that is held in a major collection.42 Accurate ITS rDNA sequences are available for more than half of the ex-type isolates (or representatives isolates of the species determined by specialists) of the species within the genus Penicillium.6,8,43,44 A phylogeny based on ITS plus the 5′ end of the lsu rDNA (1240 nucleotide) of 100 ex-type isolates covering the subgenera Aspergilloides, Furcatum, and Penicillium together with Eupenicillium species8 uncovered a more complicated intrageneric organization than proposed on the basis of phenotypic criteria.1 The subgenus Penicillium forms a consistent clade, but includes some Furcatum species in a basal position, while remaining Furcatum and Aspergilloides species are intermingled into five other different clades. The sequences used in this study have shown little genomic diversity within the subgenus Penicillium clade, in comparison to the other clades. And some important, terverticillate species are not detected when using ITS rDNA alone in a quantitative PCR analysis.44 A more rapidly evolving gene coding for β-tubulin, was used to entirely re-evaluate the 58 species accepted in the subgenus Penicillium using several isolates for each species.2 A number of species were shown to be polyphyletic and there was still some related species unresolved. COI used on the same sampling did not increase the resolution.45 Among the terverticillate species of interest in this chapter, P. chrysogenum and P. brevicompactum have been subjected to GCPSR.3,4 P. chrysogenum was shown to be composed of

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three clonal lineages well supported in the four genes phylogenies (ITS rDNA, β-tubulin, acetyl co-enzyme A synthase and thioredoxin reductase). The authors have not split this species in the way to conserve the actual nomenclature as suggested by Frisvad et al.46 for species of industrial importance. GCPSR using ITS rDNA, partial elongation factor and calmodulin genes demonstrated that P. brevicompactum, P. biourgeanum, and P. olsonii were three distinct species, diagnosable by ITS rDNA sequences. GCPSR of P. commune is in progress in our laboratory showing the presence of three distinct lineages (Dupont et al., unpublished data), that are not diagnosable by ITS rDNA. P. carneum and P. paneum, first described as varieties of P. roqueforti according to their secondary metabolite profiles,47 have been elevated to the rank of species after molecular evaluation within ITS rDNA and by RAPD.48 An intra-specific variability was observed in the important mycotoxigenic P. expansum and P. griseofulvum using the β-tubulin sequences2 and examination of other isolates is necessary to evaluate and take into account the extent of this variability for molecular diagnosis. In the same study, the delimitation between P. verrucosum and P. nordicum was problematic as P. verrucosum derived within P. nordicum, needing GCPSR assessment on more numerous isolates. P. sclerotigenum, a patulin producer, classified in the subgenus Furcatum by Pitt1 is in fact a well defined species related to P. expansum.2 Only few species of the subgenus Biverticillium and related Talaromyces have been phylogenetically investigated.6,43 A collaborative GCPSR project is in progress for the complete revision of this group and ITS rDNA is shown to be a good marker to discriminate species in this subgenus (Dupont et al., unpublished). However the regions of the β-tubulin and translation elongation faction genes that have been sequenced carry large introns that are cumbersome to use at the scale of the whole subgenus but very useful for GCPSR assessment of related species (Dupont et al, unpublished data). Recently, partial calmodulin sequences were tested for their phylogenetic value on a comparable sampling to the one used by Peterson8 on Chinese isolates representing 56 species from which several are biverticillate.49 This study brings information on interspecific relationships and global phylogenetic structure of the genus, although the main branches of the phylogentic tree published are not strongly supported. But there are little insights on the species perimeter as very few isolates were analyzed for each species. Nevertheless calmodulin, together with β-tubulin have been used successfully to describe new species.50 Besides the genes that are commonly utilized for phylogenetic analysis and molecular taxonomy of fungi, genes involved in mycotoxin production and decay of fruit have been investigated for detection of mycotoxigenic species. The polyketase synthase gene, highly induced under OTAproducing conditions in P. nordicum, which did not amplify the closely related P. verrucosum, was used to develop a realtime PCR system for the detection of P. nordicum in wheat.51 OTA biosynthesis pathway is however still hypothetical and the question about potentially false positive citrinin-producing

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fungi was raised52 as OTA is a combination of the polyketide structure basically of the mycotoxin citrinin with the amino acid phenylalanine. Species-specific primers were designed, from the isoepoxydon dehydrogenase gene involved in the patulin biosynthetic pathway (extremely specific unlike that of OTA), for the identification of Penicillium linked to patulin contamination53 on the basis of previous work attesting to the usefulness of this gene in fungal taxonomy.54,55 In the same way, primers were designed to amplify specifically the polygalacturonase gene of P. expansum which encodes for the enzyme responsible for the decay of fruit tissue56 but the authors did not include in their experiments closely related species P. marinum and P. sclerotigenum which are patulin producers.13 An important molecular work has been realized on Penicillium, but very few papers have studied a significant number of strains for each taxon, to be useful for species recognition. In each Penicillium species that have been evaluated accurately by molecular tools, several phylogenetic species are recognized. And this may be expected for a number of poorly defined species for which the taxonomy has been very confused. Presently, ITS rDNA, being assigned as the core target molecule for fungal species identification, is already available for more than half of the species of the genus. We expect that the Barcode of Life initiative will be taken up and coordinated by specialists of the genus to fill the gap very soon. However ITS alone will not resolve all the species of the genus and complementary sequences are necessary to identify some closely related species. β-tubulin have significantly improved the resolution of terverticillate species and accurate references exist already in public databases. An extensive multigenic revision of biverticillate species is in progress and will provide accurate tools rapidly. There is an undeniable delay in the revision of subgenera Aspergilloides and Furcatum and molecular references are very partial.

42.2  Methods 42.2.1 Samples Preparation 42.2.1.1  Isolation of Penicillium from the Air To detect Penicillium from the air, an isolation step is required as many fungal spores are present in the airspora. The simplest method of air sampling is by sedimentation on settle plates, containing an appropriate agar isolation medium. However volumetric air sampling is more reliable. A number of air sampling devices are commercially available. The use of different media is recommended to have a better chance to isolate suspected Penicillium. dichloran rose bengal chloramphenicol agar (DRBC) contains growth inhibitors (dichloran and rose bengal) which restrict colony spreading, particularly of fast growing fungi like Rhizopus and Mucor, in the way to obtain isolated colonies that are easy to count and to pick for subculturing. Dichloran 18% glycerol agar (DG18) is useful to isolate xerophilic species from low moisture foods such as stored grains, nuts, flour, and spices. MEA with chloramphenicol (MAChl) is a standard isolation medium

Molecular Detection of Foodborne Pathogens

used in mycology. Penicillium colonies, identified under the stereomicroscope will be subcultured for their maintenance and further studies on the standard medium malt agar (MA). From air conditioners, colonies may develop inside the system and be treated as on food substrates. 42.2.1.2  Isolation of Penicillium from Food Samples When contamination by Penicillium is visible on the food sample (apples, grapes, dairy, and meat products, etc.) and when only one species is suspected, a few spores may be picked directly from the colony to inoculate a single point (streaking techniques are ineffective for filamentous fungi and are not recommended) on a MA plate. As Penicillium produce highly similar colonies, several species may be present. In this case, serial spores dilutions in aqueous 0.1% peptone (the addition of a wetting agent such as polysorbitan 80 may be desirable but the natural ability of the peptone is usually adequate) may be used to isolate colonies and differentiate potential different species by eye or under the stereomicroscope. Cereals, beans, and nuts contamination is sometimes internal and it is better to blend the sample and to use dilution methods to reveal Penicillium colonies. To check for the presence of Penicillium spores in powders like flour and spices, a serial dilution method is also recommended. 42.2.1.3 Preparation of Fungal Cultures for Toxin Detection To test if isolated Penicillium from food matrix can produce mycotoxins, four efficient culture media for mycotoxin production are CYA, yeast extract sucrose agar (YES), rice powder corn steep agar (RC) and Mercks MEA (MME).16 Potato dextrose broth supplemented with manganese (152 μmol/l) was recommended for patulin production by Penicillium.57 The preparation of samples to be used in the standardized liquid chromatography–UV-mass spectrometry microscale method for screening of fungal metabolites and mycotoxins in culture extracts58 is as follows: 6 mm plugs are extracted twice in a 2-ml vial for 30 min, first using 1 ml ethyl acetate with 1% formic acid and then using 1 ml isopropanol; the pooled extracts are evaporated in a rotational vacuum concentrator. Residue is dissolved in 400 μl methanol, ultrasonicated for 10 min, and filtered through a 0.45-μl PTFE syringe filter. 42.2.1.4 DNA Extraction from Fungal Cultures for Molecular Identification DNA is extracted from fresh mycelium grown on MA plates for 24–48 h. Several kits work well, e.g., the DNeasy Plant mini kit (Qiagen). Mycelium, necessary to fill up a third of a 2-ml microtube, is scraped from the surface of the MA plate. DNA obtained are diluted 1/10 in ultra pure water for PCR amplification. Alternatively, DNA can be isolated using a method adapted from Möller et al.59 using CTAB (cetyltrimethylammoniumbromide) for efficient removal of polysaccharides highly produced by some Penicillium species. Briefly,

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Penicillium

mycelia (24–48 h old) are harvested from the surface of a MA plate and grinded to a powder in liquid nitrogen with a mortar and pestle. The mycelia powder is resuspended in lysis buffer (50 mM Tris, pH 8.0, 100 mM NaCl, 5 mM EDTA, 1% SDS). Add 50 μg proteinase K, and incubate 30 min (minimum) at 60°C. Add salt concentration to 1.4 M with 5 M NaCl and 1/10 vol 10% CTAB, incubate for 10 min at 65°C. Add 1 vol SEVAG (chloroform:isoamylalcohol, 24:1, v/v), incubate for 30 min at 0°C and centrifuge for 10 min at 4°C, 13,000 rpm. Transfer supernatant to a fresh tube; add 200 μl 3 M sodium acetate (pH 5.2), mix gently, place on ice for 30 min (minimum). Centrifuge for 10 min at 4°C, transfer supernatant to a fresh tube and add 0.55 vol isopropanol to precipitate DNA; centrifuge immediately for 5 min, 13,000 rpm. If no DNA lumps become visible immediately place samples on ice before centrifugation for approx. 15–30 min. Aspirate off supernatant and wash pellet twice with cold 70% ethanol, dry pellet and dissolve it in 50–10 μl TE buffer. A bead-agitation method has been tested56 to rapidly extract DNA from spores (4–5 h) instead of the conventional methods using fresh mycelium recovered from culture (48 h).

42.2.2  Detection Procedures 42.2.2.1  PCR Detection of Penicillium Species Principle: The ITS offer valuable targets for molecular identification of Penicillium. White et al.60 designed a pair of primers (i.e., ITS4: 5′-TCCGTAGGTGAACCTGCGG-3′; and ITS5: 5′-TCCTCCGCTTATTTGATATGC-3′) that amplify a fragment of approximately 600 bp across the ITS1-5.8S-ITS2 rDNA region, facilitating identification of Penicillium subgenus Penicillium species. Similarly, Pedersen et al 61 developed a PCR assay targeting the 5.8S-ITS2 rDNA region with primers ITS212d (5′-AAATATAAATTATTTAAAACTTTC-3′) and ITS549 (5′-CTGGATAAAAATTTGGGTTG-3′), which result in the amplification of a 336-bp fragment from Peni­ cillium subgenus Penicillium species. Alternatively, other Penicillium genes can be utilized for diagnostic purpose. For example, Glass, and Donaldson62 described the application of primers Bt2a (5′-GGTAACCAAATCGGTGCTGCTTTC-3′) and Bt2B (5′-ACCCTCAGTGTAGTGACCCTTGGC-3′) for specific amplification of a 410-bp fragment from the 5′ end of the β-tubulin gene. The PCR procedure below is based on that described by White et al.60 Procedure: (1) Prepare PCR mixture (25 μl) containing 2.5 μl of 10 × PCR buffer, 1 μl of 5 mM dNTPs, 0.625 U of AmpliTaq DNA polymerase (QBIOgène, Illkirsh, France), 1 μl of each 10 μM of each primer (ITS4 and ITS5), 50 ng of template DNA and deionized H2O (to 25 μl). Include a tube containing PCR mix but not template DNA as negative control. (2) Perform PCR amplification in a thermocycler (e.g., GenAmp 9700) for one cycle of 94°C for 2 min;

30 cycles of 94°C for 30 sec, 55°C for 30 sec and 72°C for 40 sec; one cycle of 72°C for 5 min. (3) Separate the PCR products on a 1% agarose gel containing 0.5 μg/ml ethidium bromide and visualize with a UV trans-illuminator. Note: A fragment of approximately 600 bp is amplified from Penicillium subgenus Biverticillium species.63 42.2.2.2 PCR Detection of Ochratoxin A (OTA)-Producing Penicillium Principle: Being a secondary metabolite produced by several species of Penicillium (e.g., P. verrucosum and P. nordicum) and Aspergillus (eg, A. carbonarius) moulds, OTA is a polyketide mycotoxin with nephrotoxic, hepatotoxic, immunogenic, and teratogenic properties. Given its presence in a variety of foods including cereals, beans, groundnuts, spices, dried fruits, grapes, wine, beer, coffee, and cocoa, etc, OTA poses considerable health risk to humans as a foodborne toxic substance. Prior to the development of molecular techniques, the widely utilized analytic method for OTA is liquid chromatography (LC) coupled with fluorescence detection (FLD). P. verrucosum and P. nordicum are known OTAproducing Penicillium species, with P. verrucosum being present in cereals and cereal products with P. nordicum dominating in proteinaceous foods (e.g., cheeses and fermented meats). Following the recent analysis of Penicillium genes encoding the OTA polyketide synthase (otapksPN) (responsible for the biosynthesis of the polyketide part) and the nonribosomal peptide synthetase (otanpsPN) (responsible for the linkage of the phenylalanine moiety to the polyketide),64 it becomes clear that both P. verrucosum and P. nordicum possess an identical otanpsPN gene, while they differ in their polyketide ­synthase gene. This has led to the development of a PCR assay for identification and differentiation of P. nordicum and P. verrucosum that utilizes primers (otapks_ for: 5′-tacggccatcttgagcaacggcactgcc-3′ and otapks_rev: 5′-atgcctttctgggtccagta-3′) for amplification of a 750-bp fragment from the otapksPN gene, and primers (otanps_for: 5′-agtcttcgctgggtgcttcc-3′ and otanps_rev: 5′-cagcacttttccctccatctatcc-3′) for amplification of the otanpsPN gene.65 Since the otanpsPN gene is present in both ochratoxigenic P. nordicum and P.  verrucosum, and the otapksPN gene is present only in P.  nordicum, these two OTA-producing penicillia can be rapidly detected and differentiated.65 Procedure: (1) Prepare PCR mixture (50 μl) containing 5 μl of 10 × PCR buffer, 8 μl of each dNTP (2.5 mM each dATP, dCTP, dGTP, and dTTP), 1.25 μl (5 pM) of each primer (otapks_for, otapks_rev, otanps_for, and otanps_rev), 0.5 μl (5 U/μl) of Taq polymerase (Amersham-Pharmacia, Uppsala, Sweden), 5.0 μl (0.1 μg/ml) of DNA, and 29.0 μl of deionized H2O.

600

Include a tube containing PCR mix but not template DNA as negative control. (2) Perform PCR amplification in a Biorad iCycler (Hercules, USA) for 33 cycles of 95°C for 30 sec, 60°C for 40 sec and 72°C for 60 sec. (3) Separate the PCR products on a 1% agarose gel containing 0.5 μg/ml ethidium bromide and visualize with a UV trans-illuminator. Note: The expected PCR products are about 750 bp for the otanpsPN primers and 500 bp for the otapksPN primers. P. nordicum is effectively differentiated from P. verrucosum by showing only a specific fragment of 500 bp from the otapksPN gene instead of two fragments of 750 and 500 bp from both otanpsPN and otapksPN genes as in the case of P. verrucosum.65 The specificity of the PCR assay was confirmed by its negative reaction with various food related filamentous fungi including OTA-producing Aspergillus species, suggesting the underlying genetic differences in the OTA biosynthesis in Penicillium and Aspergillus. This method was successfully applied to the analysis of the natural occurrence of P. nordicum in cured meat and ripening rooms. 42.2.2.3 PCR Detection of Patulin-Producing Penicillium Principle: A variety of fungi belonging to the ascomycete genera Byssochlamys and Eupenicillium produce patulin, which is a tetraketide mycotoxin. Apart from several patulin-producing Aspergillus species (e.g., A. clavatus, A. giganteus, A. terreus), P. carneum, Paecilomyces variotii and P. glandicola produce this toxin in silage, while P. coprobium, P. glandicola, P. vulpinum, P. clavigerum and P. concentricum are dominant patulin-producers among coprophilic fungi. Of the Penicillium species related to food products, P. expansum and P. griseofulvum have been shown to secrete patulin in apples and unfermented apple juice, and thus they represent the main source of the toxin in human consumption and foodborne illness.47 Because the gene coding for isoepoxidon dehydrogenase (idh) is involved in patulin biosynthesis, PCR primers (i.e., IDH1: 5′-CAATGTGTCGTACTGTGCCC-3′; and IDH2: 5′-ACCTTCAGTCGCTGTTCCTC-3′, yielding a 620-bp fragment from patulin-producing Penicillium strains, but 400 bp and/or 500 bp fragment from nonpatulin-producing Penicillium strains), have been designed from this gene for specific identification of patulin-producing Penicillium and other fungi.66 While the idh is widespread in the genus Penicillium, it is also found in a small number of Aspergillus, Paecilomyces, and Byssochlamys strains (which tend show fragment of 450 bp).55 Procedure: (1) Prepare PCR mixture (50 μl) containing 5 μl of 10 × PCR buffer, 6 μl (25 mM) of MgCl2, 4 μl of each dNTP (2.5 mM each dATP, dCTP, dGTP, and dTTP), 1 μl (50 pmol) of each primer (IDH1

Molecular Detection of Foodborne Pathogens

and IDH2), 0.5 μl (5 U/μl) of Taq polymerase (Amersham-Pharmacia, Uppsala, Sweden), 2.5 μl (0.1 μg/ml) of DNA, and deionized H2O (to 50 μl). Include a tube containing PCR mix but not template DNA as negative control. (2) Perform PCR amplification in a thermocycler (e.g., Hybaid) for one cycle of 94°C for 2 min; 30 cycles of 94°C for 1 min, 52°C for 1 min and 72°C for 1 min; and one cycle of 72°C for 2 min. (3) Load 10 μl of the PCR products on a 1% agarose gel containing 0.5 μg/ml ethidium bromide in TAE buffer together with a DNA molecular weight marker in a spare lane, electrophorese at 5 V/cm for 60 min, and visualize with a UV trans-illuminator. Note: A 620-bp product is generated with primers IDH1 and IDH2 from patulin-producing Penicillium strains, fragments at 400 bp and/or 500 bp are obtained from nonpatulin producing Penicillium strains, and a 450-bp fragment is observed from some Aspergillus and Byssochlamys nivea strains.54,55 Therefore, in combination with a specific PCR assay based on ITS gene sequence for Penicillium (as outlined in Section 42.2.2.1), application of primers IDH1 and IDH2 is valuable to pinpoint Penicillium isolates that produce the harmful patulin mycotoxin.

42.3 Conclusions and Future Perspectives Penicillium comprise a diverse group of filamentous fungi that may amount to over 500 distinct species. As secondary metabolites, Penicillium produce at least 18 mycotoxins, of which OTA and patulin are commonly associated with foodborne illness in humans. For this reason, it is important to identify and detect Penicillium species that produce harmful mycotoxins. Traditionally, Penicillium species are diagnosed on the basis of their morphological and biochemical characteristics involving microscopic examination. The confirmation of Penicillium mycotoxins such as OTA and patulin is achieved with LC. Considering the morphological similarities among Penicillium species and the time-consuming and variable nature of the phenotypic procedures, the use of molecular techniques for improved discrimination of Penicillium species and toxigenic strains is not only important but also necessary. Although a number of conserved gene regions (e.g., ITS and β-tubulin) have been exploited for specific identification of Penicillium subgenera Penicillium species, there is a need to identify additional genes with potential for genusspecific determination of Penicillium. Subsequent analysis of the nucleotide sequences from these genes may also help clarify the phylogenetic relationships among Penicillium species. The recent development of PCR assays for OTAproducing P. nordicum and P. verrucosum and patulinproducing Penicillium species has greatly enhanced our capability to monitor Penicillium implicated in human foodborne malaises. Future modification of these assays

Penicillium

into multiplex and real-time formats will provide a timely and ease of performance tool for food and feed quality control purposes.

References















1. Pitt, J.I. The Genus Penicillium and Its Teleomorphic States Eupenicillium and Talaromyces. Academic Press, London, 1979. 2. Samson, R.A. and Frisvad, J.C. Penicillium subgenus Penicillium: new taxonomic schemes, mycotoxins and other extrolites. Stud. Mycol., 49, 2004. 3. Scott, J., et al. Genotyping variation in Penicillium chrysogenum from indoor environments. Mycologia, 96, 1095, 2004. 4. Peterson, S.W. Multilocus DNA sequence analysis shows that Penicillium biourgeanum is a distinct species closely related to P. brevicompactum and P. olsonii. Mycol. Res., 108, 434, 2004. 5. Pitt, J.I., Samson, R.A. and Frisvad, J.C. List of accepted species and their synonyms in the family Trichocomaceae. In Integration of Modern Methods for Penicillium and Aspergillus Classification, pp. 9–49. Samson S.A. and Pitt J.I. (Eds.). Harwood Academic Publishers, Amsterdam, 2000. 6. LoBuglio, K.F., Pitt, J. and Taylor, J.W. Phylogenetic analysis of two ribosomal DNA regions indicates multiple independent losses of a sexual Talaromyces state among asexual Penicillium species in subgenus Biverticillium. Mycologia, 85, 592, 1993. 7. Peterson, S.W. Molecular genetic assessment of relatedness of Penicillium subgenus Penicillium. In The Fungal Holomorph: Mitotic, Meiotic and Pleomorphic Speciation in Fungal Systematics, pp. 121–128. Reynolds, D.R. and Taylor, J.W. (Eds.). CAB International, Wallingford, 1993. 8. Peterson, S.W. Phylogenetic analysis of Penicillium species based on ITS and LSU-rDNA nucleotide sequences. In Integration of Modern Taxonomic Methods for Penicillium and Aspergillus Classification, pp. 163–178. Samson, R.A. and Pitt, J.I. (Eds.). Harwood Academic Publishers, Amsterdam, 2000. 9. Berbee, M.L. et al. Is Penicillium monophyletic? An evaluation of phylogeny in the family Trichocomaceae from 18S, 5.8S and ITS ribosomal DNA sequence data. Mycologia, 87, 210, 1995. 10. Pitt, J.I. and Hocking A. Fungi and Food spoilage, 2nd ed. Aspen Publishers, Inc., Gaithersburg, 1999. 11. Samson, R.A. et al. Introduction to Food- and Airborne Fungi, 6th ed. Centraalbureau voor schimmelcultures, Utrecht, 2002. 12. Andersen, B. and Thrane U. Foodborne fungi in fruit and cereals and their production of mycotoxins. In Advances in Food Mycology, pp. 137–152. Hocking, A.D. et al. (Eds.). Springer, New York, 2006. 13. Frisvad J.C. et al. Mycotoxins, drugs and other extrolites produced by species in Penicillium subgenus Penicillium. Stud. Mycol., 49, 201, 2004. 14. Teubler, M. and Engels G. Low risk of mycotoxin production in cheese. Microbiol. Aliments Nutri., 1, 193, 1983. 15. EFSA (European Food Safety Authority). Opinion of the Scientific Panel on contaminants in the food chain on a request from the Commission related to ochratoxin A in food (Adopted on 4 April 2006). EFSA J., 365, 1, 2006. Available at URL: http://www.efsa.eu.int/EFSA/Scientific_Opinion/ contam_op_ej365_ochratoxin_a_food_summary_en1,0.pdf

601 16. Frisvad J.C. et al. Important mycotoxins and the fungi which produce them. In Advances in Food Mycology, pp. 3–31. Hocking, A.D. et al. (Eds.). Springer, New York, 2006. 17. Atoui A. et al. Partitioning of ochratoxin A in mycelium and conidia of Aspergillus carbonarius and the impact on toxin contamination of grapes and wine. J. Appl. Microbiol., 103, 961, 2007. 18. Boudra, H., Le Bars, P. and Le Bars, J. Thermostability of ochratoxin A in wheat under two moisture conditions. Appl. Environ. Microbiol., 61, 1156, 1995. 19. FAO/WHO (Food and Agriculture Organisation/World Health Organisation). Worldwide regulations for mycotoxins in food and feed in 2003, Van Egmond, H.P. and Jonker, M.A. In FAO Food and Nutrition Paper (FAO), no. 81 / FAO, Rome (Italy). Food and Nutrition Div., WHO, Geneva, Switzerland, 2004. 20. EC (European Community). Assessment of dietary intake of ochratoxin A by the population of EU Member States. Report of the Scientific Cooperation, Task 3.2.7. Directorate-General Health and Consumer Protection, European Commission, 2002. Available at URL: europa.eu.int/comm/food/fs/scoop/ 3.2.7_en.pdf. 21. Peraica, M. et al. The occurrence of ochratoxin A in blood in general population of Croatia. Toxicol. Lett., 110, 105, 1999. 22. Peraica, M. et al. Variations of ochratoxin A concentration in the blood of healthy populations in some Croatian cities. Arch. Toxicol., 75, 410, 2001. 23. IARC (International Agency for Research on Cancer). Ochratoxin A (Group 2B). Summaries and Evaluations, 56, 489, 1993. Lyon, France. Available at URL: http://www. inchem.org/documents/iarc/vol56/13-ochra.html 24. IARC (International Agency for Research on Cancer). Patulin (Group 3). Summaries and Evaluations, 40, 83, 1986. Lyon, France. Available at URL: http://www.inchem.org/documents/ iarc/vol40/patulin.html 25. De Hoog, G.J., Gené, J. and Figueras, M.J. Atlas of Clinical Fungi, 2nd ed. Centraalbureau voor Schimmelcultures, Universitat Rovira i Virgili, Utrecht/Reus, 2000. 26. Lyratzopoulos, G. et al. Invasive infection due to Penicillium species other than P. marneffei. J. Infect., 45, 184, 2002. 27. Vanittanakom, N. et al. Penicillium marneffei infection and recent advances in the epidemiology and molecular biology aspects. Clin. Microbiol. Rev., 19, 95, 2006. 28. Kantarcioglu, A.S. et al. Central nervous system infection due to Penicillium chrysogenum. Mycoses, 47, 242, 2004. 29. Barcus, A.L., Burdette S. and Herchline, T.E. Intestinal invasion and disseminated disease associated with Penicillium chrysogenum. Ann. Clin. Microbiol. Antimicrob., 4, 21, 2005. 30. Campbell, J. et al. Cheese worker’s hypersensibility pneumonitis. Am. Rev. Respir. Dis., 127, 495, 1983. 31. Rivero, M.G. et al. Salami worker’s lung. Medicina, 59, 367, 1999. 32. Guglielminetti, M. et al. Respiratory syndrome very similar to extrinsic allergic alveolitis due to Penicillium verrucosum in workers in a cheese factory. Mycopathologia, 149, 123, 2001. 33. Frisvad J.C., Nielsen K.F. and Samson R.A. Recommendations concerning the chronic problem of misidentification of mycotoxigenic fungi associated with foods and feeds. In Advances in Food Mycology, pp. 33–46. Hocking, A.D. et al. (Eds.). Springer, New York, 2006. 34. Paterson, R.R.M., Venancio, A. and Lima, N. Solutions to Penicillium taxonomy crucial to mycotoxin research and health. Res. Microbiol., 155, 507, 2004. 35. Taylor, J.W. et al. Phylogenetic species recognition and species concepts in fungi. Fungal Genet. Biol., 31, 21, 2000.

602 36. Hawksworth, D.L. Pandora’s mycological box: molecular sequences vs. morphology in understanding fungal relationships and biodiversity. Rev. Iberoam. Micol., 23, 127, 2006. 37. Koufopanou, V., Burt, A. and Taylor J.W. Concordance of gene genealogies reveals reproductive isolation in the pathogenic fungus Coccidioides immitis. Proc. Natl. Acad. Sci. USA, 94, 5478, 1997. 38. Geiser, D.M. et al. The phylogenetics of mycotoxin and sclerotium production in Aspegillus flavus and Aspergillus oryzae. Fungal Genet. Biol., 31, 169, 2000. 39. Geiser, D.M. et al. The current status of species recognition and identification in Aspergillus. Stud. Mycol., 59, 1, 2007. 40. Hebert, P.D.N. et al. Biological identifications through DNA barcodes. Proc. R. Soc. Lond. B, 270, 313, 2003. 41. Bridge P.D. et al. On the unreliability of published DNA sequences. New Phytol., 160, 43, 2003. 42. Ratnasingham, S. and Hebert, P.D. BOLD: The Barcode of Life Data System (www.barcodinglife.org). Mol. Ecol. Notes, 7, 335, 2007. 43. LoBuglio, K.F., Pitt, J. and Taylor, J.W. Independent origins of the synnematous Penicillium species P. duclauxii, P. clavigerum and P. vulpinum, as assessed by two ribosomal DNA regions. Mycol. Res., 98, 250, 1994. 44. Haugland, R.A. et al. Quantitative PCR analysis of selected Aspergillus, Penicillium and Paecilomyces species. System. Appl. Microbiol., 27, 198, 2004. 45. Seifert, K.A., et al. Prospects for fungus identification using CO 1 DNA barcodes, with Penicillium as a test case. Proc. Natl. Acad. Sci. USA, 104, 3091, 2007. 46. Frisvad, J.C. et al. Proposals to conserve important species names in Aspergillus and Penicillium. In Modern Concepts in Penicillium and Aspergillus Classification, pp. 83–90. Samson R.A. and Pitt, J.I. (Eds.). Plenum Press, New York, 1990. 47. Frisvad, J.C. and Filtenborg, O. Terverticillate penicillia: chemotaxonomy and mycotoxin production. Mycologia, 81, 837, 1989. 48. Boysen, M.E. et al. Reclassification of the Penicillium roqueforti group into three species on the basis of molecular genetic and biochemical profiles. Microbiology, 142, 541, 1996. 49. Wang, L. and Zhuang, W.Y. Phylogenetic analyses of penicillia based on partial calmodulin gene sequences. BioSystems, 88, 113, 2007. 50. Peterson, S.W. and Sigler, L. Four new Penicillium species having Thysanophora-like melanized conidiophores. Mycol. Res., 106, 1109, 2002. 51. Geisen, R. et al. Development of a real time PCR system for detection of Penicillium nordicum and for monitoring ochratoxin A production in foods by targeting the ochratoxin polyketide synthase gene. System. Appl. Microbiol., 27, 501, 2004. 52. Paterson, R.R.M. Identification and quantification of mycotoxigenic fungi by PCR. Process Biochem., 41, 1467, 2006.

Molecular Detection of Foodborne Pathogens 53. Dombrink-Kurtzman, M.A. and McGovern A.E. Speciesspecific identification of Penicillium linked to patulin contamination. J. Food Prot., 70, 2646, 2007. 54. Paterson, R.R., Venâncio, A. and Lima, N. A novel identification system based on 318 penicillia strains using the isoepoxydon dehydrogenase gene and patulin production. Rev. Iberoam. Micol., 23, 155, 2006. 55. Paterson, R.R.M. The isoepoxydon dehydrogenase gene PCR profile is useful in fungal taxonomy. Rev. Iberoam. Micol., 24, 289, 2007. 56. Marek, P., Annamalai, T. and Venkitanarayanan, K. Detection of Penicillium expansum by polymerase chain reaction. Int. J. Food Microbiol., 89, 139, 2003. 57. Dombrink-Kurtzman, M.A. and Blackburn J.A. Evaluation of several culture media for production of patulin by Penicillium species. Int. J. Food Microbiol., 98, 241, 2005. 58. Nielsen, K.F. and Smedsgaard, J. Fungal metabolite screening: database of 474 mycotoxins and fungal metabolites for dereplication by standardized liquid chromatography-UVmass spectrometry methodology. J. Chromatogr. A, 1002, 111, 2003. 59. Möller, E.M., et al. A simple and efficient protocol for isolation of high molecular weight DNA from filamentous fungi, fruit bodies, and infected plant tissues. Nucleic Acids Res., 20, 6115, 1992. 60. White, T.J. et al. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In PCR Protocols: A Guide to Methods and Applications, pp. 315–322. Innis, M.A. et al. (Eds.). Academic Press, San Diego, 1990. 61. Pedersen, L.H. et al. Detection of Penicillium species in complex food samples using the polymerase chain reaction. Int. J. Food Microbiol., 35, 169, 1997. 62. Glass, N.L. and Donaldson, G.C. Development of primer sets designed for use with PCR to amplify PCR conserved genes from filamentous Ascomycetes. Appl. Environ. Microbiol., 61, 1323, 1995. 63. Dupont, J. et al. PCR-RFLP of ITS rDNA for the rapid identification of Penicillium subgenus Biverticillium species. Rev. Iberoam. Micol., 23, 145, 2006. 64. Karolewiez, A. and Geisen, R. Cloning a part of the ochratoxin A biosynthetic gene cluster of Penicillium nordicum and characterization of the ochratoxin polyketide synthase gene. Syst. Appl. Microbiol., 28, 588, 2005. 65. Bogs, C., Battilani, P., and Geisen R. Development of a molecular detection and differentiation system for ochratoxin A producing Penicillium species and its application to analyse the occurrence of Penicillium nordicum in cured meats. Int. J. Food Microbiol., 107, 39, 2006. 66. Paterson, R.R.M. Primers from the isoepoxydon dehydrogenase gene of the patulin biosynthetic pathway to indicate critical control points for patulin contamination of apples. Food Control, 17, 741, 2006.

43 Rhodotorula Diego Libkind

Biodiversity and Environmental Research Institute (CONICET-UNComahue)

José Paulo Sampaio

Nova de Lisboa University

Contents 43.1 Introduction.................................................................................................................................................................... 603 43.1.1 Taxonomy of the Genus Rhodotorula............................................................................................................... 603 43.1.2 Environmental Distribution of Rhodotorula spp. and Occurrence in Food and Beverages............................. 604 43.1.3 Pathogenesis and Medical Importance of Rhodotorula................................................................................... 605 43.1.4 Limitations of Conventional Identification of Yeasts in the Clinical Laboratory............................................ 606 43.1.5 Molecular Detection of Yeast Pathogens with Special Reference to Rhodotorula spp.................................... 607 43.2 Methods.......................................................................................................................................................................... 609 43.2.1 Reagents and Equipment ................................................................................................................................. 609 44.2.2 Sample Treatment............................................................................................................................................. 609 43.2.3 Detection Procedures........................................................................................................................................ 609 43.2.3.1 Identification and Genotyping of Rhodotorula spp. by Micro/mini-satellite Primed PCR (MSP-PCR)................................................................................................................. 609 43.2.3.2 Detection of Rhodotorula spp. Employing Species-specific PCR Primers......................................610 43.2.3.3 Identification of Rhodotorula spp. by Restriction of the Amplified 5.8S-ITS Region of the Ribosomal rRNA Operon (RFLP-ITS)..................................................................................611 43.2.3.4 Identification of Rhodotorula spp. by rDNA Sequencing Analysis..................................................611 43.3 Conclusions and Future Perspectives..............................................................................................................................612 References...................................................................................................................................................................................613

43.1  Introduction When dealing with the isolation of fungi from clinical ­samples, in a few cases, yeast colonies with a characteristic salmon-pink to coral-red color appear. A classic procedure of many medical mycologists is to assign such isolates to the genus Rhodotorula on the basis of this sole phenotypic attribute. Today, thanks to the advancement of molecular biology techniques (especially DNA sequencing) and molecular phylogenetics, it is well known that yeasts producing pigmented colonies (also called pigmented or red yeasts) may belong to several basidiomycetous genera other than Rhodotorula. Examples are Rhodosporidium (teleomorphic stage of many Rhodotorula species), Sporobolomyces, Sporidiobolus, Dioszegia, Cystofilobasidium, Xanthophyllomyces, and Cryptococcus. The distinctive pink to red coloration of their colonies is the result of the intracellular production and accumulation of carotenoid compounds. The composition of carotenoid pigments may vary qualitatively in red yeasts depending on the species. The major carotenoids to date found in yeasts are torularhodin, torulene and β-carotene.1,2 However, among yeast with current medical relevancy, the pigmented species known to be most frequently involved

in human mycosis in fact belong to Rhodotorula. There are no records of clinical isolates for almost all other pigmented genera. The only exception probably is Sporobolomyces (and its sexual stage: Sporidiobolus) which has been found, in a few occasions, involved in human fungemia or have been isolated from clinical specimens.3–7

43.1.1  Taxonomy of the Genus Rhodotorula The genus Rhodotorula is polyphyletic and consists of 47 different species, constituting a very heterogeneous taxon. Most of Rhodotorula species belong to the subphylum Pucciniomycotina, either to the class Microbotryomycetes (29 species) or the class Cystobasidiomycetes (15 species). The remaining three species are classified in the subphylum Ustilaginomycotina and do not produce carotenoid pigments. R. glutinis is the type species of the genus and belongs to the order Sporidiobolales (Class Microbotryomycetes) as well as the abundant yeast R. mucilaginosa (previously known as R. rubra). A third clinically important Rhodotorula species is R. minuta, which unlike R. glutinis and R. mucilaginosa, is ascribed to the order Cystobasidiales (Class Cystobasidiomycetes). The case of R. glutinis is peculiar 603

604

Molecular Detection of Foodborne Pathogens

used to refer to R. glutinis and closely related Rhodotorula and Rhodosporidium species often confused with R. glutinis. The R. glutinis complex is hence a group of species, and its characteristics and properties correspond to that of the sum of species included in it. Figure 43.1 provides a schematic representation of the phylogenetic placement of Rhodotorula and Rhodosporidium species within the phylum Basidiomycota, as well as the main phenotypic characteristics of the species discussed in this chapter: R. mucilaginosa, R. glutinis complex, and R. minuta.

43.1.2  Environmental Distribution of Rhodotorula spp. and Occurrence in Food and Beverages Rhodotorula species can be commonly isolated from natural environments (e.g., soil, water, phylloplane, etc.). A notable case is that of R. mucilaginosa, which is one of the most (and possibly the most) ubiquitous basidiomycetous yeast species. Besides a probable association with man, Rhodotorula mucilaginosa is found in a wide range of natural habitats including living or decomposing plant constituents and soil,16–19 and various aquatic environments such as freshwater lakes,20 estuaries,21 coastal waters,22 and open ocean and deep-sea environments.23,24 Moreover, R. mucilaginosa is also present INOSITOL PAC

given that until recently it was regarded as a ubiquitous yeast and at least 370 collection strains were considered to belong to this species.8 The classical circumscription of R. glutinis was based mostly on a few physiological and cultural properties. However, several studies have reported on the physiological and genetic heterogeneity of R. glutinis.9–13 More recently, Sampaio et al.14 and Gadanho and Sampaio15 re-evaluated the circumscription of this species using a large set of isolates that had been previously identified by phenotypic criteria as R. glutinis, and found that most of the isolates actually did not belong to this species. The most common were anamorph strains of Rhodosporidium babjevae, but other species of Rhodotorula (R. dairenensis, R. graminis, and Rhodotorula spp.) and Rhodosporidium (Rh. diobovatum, Rh. sphaerocarpum) were also misidentified as R. glutinis. In such study it was shown that from a total of 45 isolates only four had been unequivocally assigned to R. glutinis. The species with the closest phenetic and phylogenetic proximity to R. glutinis are R. graminis and Rh. babjevae.15 Therefore, it can be hypothesized that most studies reporting R. glutinis isolates from human sources may have been confused with other yeast species. Furthermore, it becomes clear that the accurate identification of R. glutinis and closely related species is only possible by the use of molecular techniques. In order to avoid confusions, hereinafter the term R. glutinis complex will be

Basidiomycota

(a) Sub-phylum Agaricomycotina

Cryptococcus

Ustilaginomycotina + –

Pseudozyma / Ustilago Rhodotorula / Rhodosporidium

(b) R. mucilaginosa R. minuta

– –

R. mucilaginosa

Glucuronic acid R. minuta

+

Nitrate Gallic acid Saccharic acid +

R. glutinis

R. glutinis complex

Rhodotorula Rhodosporidium

Pucciniomycotina

– –

+ +

Rh. kratochvilovae Rh. diobovatum Rh. babjevae

– + + –

– – + +

Figure 43.1  Schematic representation of the phylogenetic placement of Rhodotorula (R., asexual) and Rhodosporidium (Rh., sexual) within the phylum Basidiomycota (a) and the main phenotypic characteristics of the species discussed in this chapter. (b) R. glutinis complex, R. minuta and R. mucilaginosa. Most species in Cryptococcus form white colonies (open circle) and a minority of species form pink colonies (grey circle). Colony colors are similar in Rhodotorula but their proportions are symmetrical. The members of the Ustilaginomycotina do not form pink colonies. The ability to utilize inositol as sole carbon source and to produce amyloid compounds (PAC) allows the discrimination of the yeasts belonging to the three main lineages of the Basidiomycota. The most relevant Rhodotorula and Rhodosporidium species can be distinguished based on the ability to utilize nitrate as sole nitrogen source and on the capacity to utilize glucuronic, gallic, and saccharic acids.

605

Rhodotorula

in extreme environments like hyper-acidic waters,25,26 uranium leachate27 and cold environments28–30 among others. The R. glutinis complex is also worldwide in distribution, and has been isolated from a variety of substrates. Known sources include: air, freshwaters, marine waters, terrestrial environments (soil, phylloplane, flowers, etc.), food and beverages, animals, and humans.8 In comparison to R. mucilaginosa and R. glutinis complex, R. minuta has been less frequently isolated in natural environs. This species was detected in air, seawater (including deep-sea environments)23 and freshwater.20 Concerning the presence of these Rhodotorula species in food, as for natural environments, R. mucilaginosa is the species with the largest frequency of appearance and the widest range of food and beverages from which it has been detected. Several studies reported the isolation of R. mucilaginosa from peanuts, apple cider,31 cherries,32 fresh fruits,33–35 fruit juice,36 cheese,36–38 sausages,39 edible mollusks,40 and crustaceans,41 among others. The remaining two Rhodotorula species are less frequent and normally when they are present, R. mucilaginosa is also present. However, a few exceptions are known, as an example Callon et al.42 reported the isolation of R. glutinis complex and R. minuta as the only representatives of pigmented yeasts from goat milk. Although consumption of food contaminated with yeasts may not have a direct role in causing opportunistic infections, there is increasing concern that foods could be an underestimated environmental source of these yeast pathogens.43 In fact, Tomsikova et al.44 studied fungal contamination of food distributed in hospitals to inmunocompromised patients and found that yeasts (including Rhodotorula spp.) were present in 82% of fruit, 13% of cheeses, and 46% of smoked meat products. Thus, it is becoming clearer that foods with substantial yeast loads (e.g., processed meats, soft cheeses, fresh fruits) pose a risk to susceptible hospital patients.45–47 In order to clarify the role of foods in the contribution to opportunistic yeast infections, further research is needed to understand (i) the survival and growth of foodborne yeasts throughout the gastrointestinal system, (ii) the potential for such yeasts to transfer from the gastrointestinal tract to the blood stream, and (iii) the general occurrence and ecology of these yeasts in hospital and health-care environments.43 Concerning the first point, Rhodotorula spp. have been isolated from stool samples,48–49 an indication that these yeasts can survive extreme conditions of the gastrointestinal tract. However, it has not been reported as a prevalent yeast of human intestinal flora.50 If these yeasts are able to colonize and grow in the digestive tract is still uncertain and needs further investigation, as well as their ability to pass to the blood stream. Our knowledge on the occurrence and ecology of Rhodotorula spp. in the hospital environment is larger than that for the first two issues. Rhodotorula species can be found in association with humans, having been isolated from various types of human specimens like feces, skin, sputum, forming part of the normal flora.51 These yeasts are generally considered commensal saprobes,52–55 occurring most frequently in the environment than in human tissues.

It is important to know that, Rhodotorula spp. have been found to be one of the most frequent isolated microorganism from nurses and nonnursing hospital personnel hands.56,57 Moreover, many patients studied on the very day of hospital admission have been shown to be carriers of such yeasts.58 Rhodotorula yeasts possess a strong affinity for plastic, having been recovered from various medical equipments like hemodialysis machines and fibreoptic bronchoscopes,59–60 and several environmental sources including shower curtains, bathtub grouts, and toothbrushes.61 A direct consequence of such broad exposure to these yeasts, is that in humans that for some reason have their immunological system depressed, Rhodotorula spp. can occasionally behave as opportunistic pathogens causing a variety of systemic infections. As a result, patients may suffer increased morbidity and mortality through colonization, allergy, and invasive infection from such yeasts.

43.1.3  P  athogenesis and Medical Importance of Rhodotorula Rhodotorula species that are regarded as emerging yeast pathogens are R. mucilaginosa (synonym R. rubra),62,63 which seems to be frequently associated with human infections, and the less frequent R. glutinis complex and R. minuta.64 Such species have been implicated in a broad spectrum of fungemia and other malignancies mostly in immunocompromised hosts, however, a few cases of Rhodotorulosis in inmuncompetent patients have been reported.65,66 To the best of our knowledge one of the first cases of fungemia caused by Rhodotorula spp. was reported in 1960 for a patient with endocarditis.67 A dramatic increase in the number of cases of Rhodotorulosis has been observed in the following years. Factors predisposing to such opportunistic infections include underlying immunosuppression, the use of broad-spectrum antibiotics and/or narrow-spectrum antifungals, and the presence of foreign devices such as central venous catheters.61,68 Though, the use of indwelling devices is by far the major risk factor for infection with these opportunistic yeasts.69–71 Rhodotorula spp. have been incriminated in cases of ocular infections,72–76 meningitis,77–80 peritionitis,81 endocarditis,82 ventriculitis,83 cancer-associated infections,53,63,84 infections related to central venous catheters, and other indwelling devices,53,55,59,63,71,79,85–88 and various others systemic infections.89–95 R. mucilaginosa is the species responsible for the majority of these medical cases (~70%) followed first by R. glutinis complex (~10%) and then by R. minuta (~6%). Reports in which Rhodotorula isolates were or could no be properly identified to the species level correspond to the remaining percentage of the cases. However, it should be noted that in many of these cases, sufficient evidence supporting the involvement of the yeast isolate in the pathogenic process is lacking. In some cases, the basis for considering Rhodotorula yeasts as pathogenic is based on resolution of fever accompanied by disappearance of the yeast from blood cultures without evidence of clearance of an infectious focus. Such evidence is suggestive that

606

the blood isolate was the etiologic agent of fever, but it is not definitive.62 With respect to other pathogenic yeasts, like those of the genus Candida, the incidence of Rhodotorula infections is quite low. Early epidemiological studies in USA and Europe showed that the incidence of Rhodotorula infections was between 0.5 and 2.3% of total fungemia cases.68,96,97 Moreover, Lunardi et al.,64 reported that in a 3-year surveillance of the occurrence of rhodotorulosis in a Brazilian tertiary care hospital, the overall incidence of such infections was 0.056 episodes per 1000 hospital admissions. In a more recent work, Pfaller et al.98 observed that from more than 200000 clinical yeast isolates obtained during an 8 year survey, Rhodotorula spp. represented the 4.2%. However, it should be noted that infections caused by unusual yeasts like Rhodotorula spp. could be under-reported because of difficulties in accurate diagnosis and a tendency of attributing isolates to specimen contamination.99 Today, Rhodotorula spp. are still rare in the diagnostic mycological laboratory,100 but an increase in the frequency of invasive mycoses due to these opportunistic fungi and many others can be anticipated. This will most probably occur due to the ever increasing application of newer technologies and therapies in medical practice resulting in longer and more severe immunosuppression of nosocomial patients. Furthermore, given the Rhodotorula spp. intrinsic resistance to the widely used triazole and echinocandin antifungal agents, patients receiving such drugs, which select for Rhodotorula spp.,65 will be more susceptible to the development of Rhodotorula fungemia.64,101 Even though Rhodotorula species appear to be less ­virulent than more common yeast pathogens like Candida and Cryptococcus neoformans for example, fatal cases in which Rhodotorula spp. was suspected to be the etiological agent have been reported. In the literature, the overall mortality rate of Rhodotorula fungemia of reported cases is around 15%,64,87,102,103 although a few authors consider it higher.69,104–106 Rhodotorula-related sepsis syndrome and other life-threatening complications have also been reported.53,70,107 The generally low pathogenicity of Rhodotorula spp. is probably related to its reduced ability to grow at 37°C, an attribute typically enhancing virulence of pathogenic strains.108 Several clinical and environmental isolates of R. glutinis complex have been reported to grow at 37°C,109 however, it should be noted that the species Rh. toruloides often misidentified as R. glutinis, and hence included in the R. glutinis complex, can grow at 37°C.8 Other explanations probably derive from the scarce ability of these yeasts to secrete degradative enzymes such as proteinase110 and phospholipases,111 which may be associated with pathogenic processes caused by opportunistic yeasts. Rhodotorula spp. infections have been successfully treated with catheter removal alone (for indwelling ­devices-associated cases), antifungal therapy and with a combination of these two approaches.64,70,112 Antifungal treatment of Rhodotorula infections typically involve the use of amphotericin B, given that Rhodotorula spp. was shown to

Molecular Detection of Foodborne Pathogens

be highly susceptible to such compound in vitro76,112 and that it has been proven to be effective for clearing various types of Rhodotorula fungeamia.70,83,113–115 Liposomal amphotericin B should be also considered because it is less nephrotoxic than conventional amphotericin B deoxycholate while maintaining antifungal activity against Rhodotorula spp.116 More than a few cases have been reported in which treatment with this antifungal worked successfully for curing various types of Rhodotorula infections.59,91,101,117 Alternative antifungals for the treatment of Rhodotorulosis include Flucytosine78,114 showing excellent in vitro activity86,118 and the promising new and investigational triazole Ravuconazole.112 Of worth noting is the evidence provided by the work of Diekema et al.112 who tested the antifungic susceptibility of 64 clinical Rhodotorula isolates including R. mucilaginosa (24 isolates), R. glutinis complex (29 isolates), R. minuta (five isolates) as well as other nonidentified Rhodotorula species (six isolates), finding no significant differences among them in the activities of any of the tested agents. These results suggest that the frequent infection-causing Rhodotorula species behave similarly toward most antifungal treatments. Interestingly, Silici et al.119 showed that six different honey bee propolis were highly effective in inhibiting a strain of Rhodotorula spp. displaying very low MIC values ( ≤ 0.01 µg/ml). Such results are promising and should be corroborated with a larger set of isolates in order to evaluate propolis as an alternative antifungal agent for Rhodotorula mycoses. Narrow spectrum azoles (e.g., fluconazole) and echinocandins (e.g., caspofungin) type antifungics should not be considered appropriate for treatment of such cases due to Rhodotorula spp. high resistance to these kind of drugs.86,112,120

43.1.4  Limitations of Conventional Identification of Yeasts in the Clinical Laboratory Conventional identification of yeast species and strains is based on morphological, physiological, and biochemical characteristics. In order to reliably identify yeast to the species level it is necessary to conduct 50–100 phenotypic tests. This procedure is laborious, time-consuming (it usually takes 3 weeks or more), and only an experienced person can obtain proper interpretation of the test results.121 Furthermore, sometimes ambiguous identifications may be obtained due to strain variability. For the identification of mycoses-causing yeasts, as is the case of some Rhodotorula species, a reliable identification is required for epidemiological investigations and for the selection of an appropriate antifungal treatment. Conventional identification of yeast isolates in the clinical laboratory is based on one of several commercially available biochemical and enzymatic panels. However, some disadvantages of these identification kits include the existence of limited databases122,123 and frequent occurrence of misidentification.87,122,124–129 Because of the ever expanding spectrum of fungal pathogens (including yeasts), the field of medical mycology has become an extremely challenging study of infections caused by opportunistic fungi. Given that

607

Rhodotorula

the identification of such wide and taxonomically diverse array of fungal pathogens by conventional methods is often difficult and sometimes inconclusive,130 molecular biology techniques are being incorporated in clinical laboratories to provide more accurate and prompt diagnosis.

43.1.5  Molecular Detection of Yeast Pathogens with Special Reference to Rhodotorula spp. Molecular diagnostic methods possess several advantages over conventional methods based on phenotypic characteristics, such as higher reliability, reproducibility, and throughput. Due to their high specificity and sensitivity, these molecular methods should be used routinely in the clinical laboratories as a complement of the information provided by conventional methods and particularly, to help in the diagnosis of dubious cases. A variety of molecular approaches for genotyping and identification of yeast isolates have been described. These methods include species-specific DNA primers,126,131–133 multiplex PCR,134,135 real-time PCR assays,135,136 PCR fingerprinting (i.e., RAPDS, MSP-PCR),14,15,20,22,137 PCR-enzyme immunoassays,138 single-strand conformation polymorphism (SSCP) analysis of ribosomal DNA,139 restriction fragment

length polymorphism (RFLP) analysis of 5.8S internal transcribed spacer (ITS) ribosomal DNA (rDNA),140-142 electrophoretic karyotyping by pulsed-field gel electrophoresis,143–145 denaturing/temperature gradient gel electrophoresis (DGGE/ TGGE),146–151 amplified fragment length polymorphism,148,152 flow cytometry or Luminex®,153–156 DNA microarray,156–158 and DNA sequencing.159,160 However, not all these techniques can be readily applied in the clinical laboratory due to their complexity, high costs and/or limitations in the array of yeast species that can detect. Several of such methods have been only developed for usual pathogenic yeasts (Candida spp., Malassezia spp., Cryptococcus neoformans) while are yet not applicable for less common opportunistic yeasts like Rhodotorula spp. For a list of available molecular techniques for yeast identification, as well as their advantages and disadvantages see Table 43.1. PCR-based techniques in general provide the required simplicity, specificity, and sensitivity to identify most fungal species in a short time. Most of these methods do not directly provide identification because this can only be achieved by comparing results with databases or running tests together with reference strains (i.e., PCR fingerprinting, multiplex PCR, ITS sequence length polymorphism, and SSCP-ITS). A PCR fingerprinting method that proved to be very useful

Table 43.1 Molecular Detection Techniques* for Yeasts with Special Reference to Rhodotorula spp. and their Characteristics Method PFGE PCR fingerprinting MSP-PCR RAPD Species specific primers/multiplex PCR Sequence length polymorphism (ITS1 or ITS2) SSCP-ITS RFLP-ITS PCR – T/DGGE D1/D2 rDNA sequence ITS rDNA sequence Other gene sequences (IGS, 18S, cytochtome b) Real-time PCR FISH* Pyrosequencing® DNA microarray Flow cytometry, Luminex®* * **

Characteristics Laborious, time consuming, lack of databases, needs reference strains, requires specific equipment. Fast, simple, needs reference strains, suitable primers may vary with yeast group. High repetitiveness, high discriminatory power. Low repetitiveness. Fast, simple, needs specific primers.** Fast, simple, limited databases Fast, simple, needs reference strains, limited to known species. Simple, specificity depends on yeast group, limited database Complicated, needs experience, low repetitiveness, useful for culture-independent detection. Highly conserved, more complete database, provides accurate identifications More variable, less complete database. Useful for analyses of closely related species Limited databases. Sensitive, fast, needs specific probes** and equipment. Simple, fast, needs specific probes**. Useful only if cells are in high numbers and metabolically active. Low cost, limited to short DNA fragments. High-throughput, needs specific DNA probes**, specific equipment. High-throughput, needs specific DNA probes**, specific equipment.

References 143–145

15,137 148,183  126,131–135,184  124,125 139,185 140–142 146–151 159,164 128,160,165 186–188,191 135,136,181 182,189 190 156–158 153–155

Techniques that to the best of our knowledge have not been yet tested or developed for Rhodotorula spp. Identification. Implies that such technique is limited to known species. PFGE: pulsed field gel electrophoresis, MSP-PCR: micro/minisatellite primed-PCR, RAPD: random amplified polymorphic DNA, SSCP: single-strand conformational polymorphism, RFLP: restriction fragment length polymorphism, T/DGGE: temperature/denaturing gradient gel electrophoresis, FISH: fluorescent in situ hybridization.

608

for the rapid and simple genotyping of Rhodotorula isolates is the micro/mini-satellite primed-PCR (MSP-PCR) technique.15,20,137 It consists in the PCR amplification of genomic DNA using synthetic oligonucleotides that detect micro/minisatellite DNA. Microsatellite DNA consists of sequences repeated motifs of ca. 2–10 bp arranged in tandem at various loci, while minisatellite DNA is made up of motifs of 15–30 bp arranged in tandem. Such regions are highly polymorphic and thus provide sufficient resolution for the discrimination among yeast species. However, the discrimination power of each primer differs and it is related to the phylogenetic position of the taxa under study. After PCR reaction, MSP-PCR DNA banding patterns are obtained by agarose gels electrophoresis. These are species specific, allowing the unambiguous identification of most yeast species tested so far by comparison with profile databases or reference strains. Often, small variations in minor bands may provide differentiation up to strain level, which is useful for epidemiological studies.137,161 Several studies have successfully employed the MSP-PCR method for rapid genotyping of large sets of collection strains and/or environmental isolates which included Rhodotorula species.14,15,20,26,30 In such cases, the species R. mucilaginosa and R. minuta were easily identified by direct comparison with reference strains. Libkind et al.137 studied almost 100 environmental isolates of R. mucilaginosa and found that more than 90% of these shared similar MSP-PCR profiles than the type strain using either M13 or (GTG)5 primers, thus allowing prompt identification. On the other hand, studies within the R. glutinis complex have demonstrated that this closely related group of species shares similar MSP-PCR banding patterns with primers like M13. The discrimination among these species can be obtained with the alternative MSP-PCR primer (GAC)5.15 To the best of our knowledge, the MSP-PCR technique has not been yet tested for clinical isolates of Rhodotorula, though; Latouche et al.161 applied this technique for Candida clinical strains and found it fast, reproducible, and more costeffective than available biochemical approaches. An alternative PCR method that can be used to directly identify Rhodotorula spp. (as well as others fungal pathogens) involves the use of species-specific primers.131,132 The technique employs three primers in a PCR amplification of a ~600 bp DNA segment, consisting of two external universal (upstream and downstream) primers and one internal speciesspecific primer. Species identification requires the formation of species-specific rDNA nucleotide amplicon which is significantly smaller than the non-target segment. Multicopy genes such as those of ribosomal origin (either the D1/D2 region of the large rDNA subunit or the noncoding ITS regions of the rRNA-encoding gene) are preferred by many researchers to achieve well defined results.131,132 However, because they may generate false positive results, other researchers have looked at single-copy genes of high specificity.162 Specific rDNA signature nucleotides or sequences of the target yeast species are used for the development of the specific oligonucleotide primers. A worth noting disadvantage of this detection technique is that primers are designed based on sequences available in the GenBank database, which may not represent

Molecular Detection of Foodborne Pathogens

the whole nucleotide variability for each species. This is particularly important for Rhodotorula spp. given that R. mucilaginosa,137,145 and R. glutinis complex15 are genetically heterogeneous species. Thus, it can be anticipated that false negative results could be obtained using species-specific primers for the molecular diagnosis of Rhodotorula species. Furthermore, to the best of our knowledge currently there is only one available specific primer for Rhodotorula spp., which was designed to detect R. mucilaginosa.132 Other techniques have been developed for rapid yeast genotyping that may or may not involve PCR (Table 43.1), one of which is the restriction fragment length polymorphism of the 5.8S-ITS rDNA sequence (RFLP-ITS). RFLPITS consists of a PCR amplification of the ITS fragment and subsequent digestion with three restriction enzymes.141 The length of the ITS fragment together with the lengths of the fragments resulting from the enzymatic restriction are used for species identification employing existing RFLP-ITS databases (i.e., www.yeast-id.com). However, it may be impossible to differentiate very closely related species using this method given that sequence differences occurring outside the restriction sites cannot be detected by RFLP-ITS analysis, and thus, fragments with identical sizes do not necessarily have identical sequences.163 Furthermore, RFLP-ITS database is still quite limited for basidiomycetous yeast species, particularly for Rhodotorula spp. One major drawback of the previously described PCRbased techniques is the fact that their diagnostic capabilities are limited to the respective existing databases, which in general only include species considered pathogenic. Thus, new species becoming emergent pathogens would not be detected with these techniques until they are incorporated into each database. Currently, the molecular technique most commonly used for accurate identification of yeasts is gene sequencing, because it offers a rapid and robust method for recognizing species and resolution is not limited to closely related taxa. Yeast species identification based on DNA sequencing, have emphasized either in coding (D1/D2 variable domains of the large subunit rDNA) or noncoding (ITS) regions of the rDNA. As a result, largely complete databases of D1/D2159 and ITS160 sequences are available for molecular classification and identification of yeasts, including all known Rhodotorula species. The value of the D1/D2 region of the 26S rDNA for identifying clinically relevant yeast isolates by DNA sequencing has been clearly demonstrated.164 Sequence analysis of this region provides enough resolution for the correct discrimination of R. mucilaginosa and R. minuta, given that both species show low genetic heterogeneity in such region.137 Different is the case of R. glutinis, given that only a single base mismatch differentiates it from the closely related species R. graminis and R. babjevae.15 Thus, D1/D2 sequence analysis may be useful for identification of R. glutinis complex, but does not provide enough resolution for the discrimination of the various related species included in this group. As an example, Leaw et al.,165 were not able to differentiate R. glutinis from Rh. babjevae and R. graminis using D1/D2 sequence analysis.

609

Rhodotorula

A higher discriminatory level may be achieved when using the more variable ITS region in rDNA sequencing studies. This sequence is equally suitable than the D1/D2 region for accurate identification of both R. minuta and R. mucilaginosa. Furthermore, it has been demonstrated that even sequencing of either the ITS1 region alone or the ITS2 region alone may be enough for the correct identification of these two yeast species.165 For the particular case of R. mucilaginosa, strain variability can also be detected with ITS sequencing,137 which is interesting for epidemiological investigations. Again, difficulties may appear when trying to discriminate species of the R. glutinis complex by sequencing the ITS region. Gadanho and Sampaio,15 using the complete ITS region, determined that R. glutinis and R. graminis differed in one mismatch and three indels, whereas three mismatches differentiated R. glutinis from R. babjevae. Studies testing the usefulness of ITS1 or ITS2 sequences alone for yeast identification, found that a strain suspected to be R. glutinis (strain BCRC 20576) could not be differentiated from Rh. babjevae using such regions.165 However, in such case, the criterion used to assign the strain BCRC 20576 to R. glutinis is not provided and thus is probable that this strain did never actually belong to R. glutinis. A key factor determining the correct identification of the sequenced yeast isolate using the BLAST tool of the GenBank, is the sequences used for comparison. Unfortunately, today the GenBank database includes many rDNA sequences that have been wrongly submitted or have been submitted with inappropriate species designations. Hence, inexperienced clinical mycologists might be leaded to incorrect identifications. When using rDNA sequencing for yeast identification one should be aware of this fact and select for comparison purposes only trustworthy sequence submissions, preferably those corresponding to type strains. Additional hints for a correct use of GenBank database while identifying Rhodotorula yeasts are given in Section 43.2.3.5.

43.2  Methods There are several molecular methods that allow the accurate differentiation and detection of Rhodotorula spp. species. In this section we provide our recommendations for molecular techniques that are appropriate for detection and genotyping of Rhodotorula isolates from clinical samples.

43.2.1 Reagents and Equipment The list of the reagents and equipment needed for the analysis relative to the sample treatment and subsequent identification of Rhodotorula spp. that will be described in detail in the following sections is depicted in Table 43.2.

44.2.2 Sample Treatment Sample collection and preparation. The sample collection and analysis procedures vary for each material drawn. With the exception of blood or swab samples which are used to inoculate appropriate liquid medium for yeast growth, other specimens are aseptically collected, homogenized,

diluted, and submitted to plating procedures. In the clinical laboratory, the isolation of yeasts is commonly performed on Sabouraud dextrose agar (10 g/l mycological peptone, 40 g/l dextrose, pH 5.6) containing 100 mg/l chloramphenicol, though Rhodotorula species may be more effectively grown in YM agar (3 g/l yeast extract, 3 g/l malt extract, 5 g/l mycological peptone, 10 g/l dextrose, pH 5). It should be point out that the Pagano Levin Medium was not suitable for the detection of R. mucilaginosa in various human specimens.49 The characterization of the yeast isolate by genetic methods should be performed only after the possibility that the strain under study is a saprophytic contaminant has been discarded. DNA extraction. Firstly, for DNA extraction, pure cultures of the Rhodotorula isolate must be obtained. For this, single colonies should be picked from the isolation plates and streaked on the surface of yeast-specific agar plates. After growth, a well isolated colony is transferred to fresh culture medium either in tubes or plates. At the moment of DNA extraction, the yeast is inoculated in the surface of fresh agar medium plates and incubated 48–72 h at room temperature (20–25°C). A rapid and simple DNA extraction protocol suitable for any of the above mentioned PCR based methods, has been thoroughly described by Sampaio et al.14 Here we provide a shorter and equally effective method for extraction of genomic DNA from a pure culture of Rhodotorula yeasts. The required reagents, consumables, and equipment are listed in Table 43.2. The procedure includes the following steps: (1) Grow the Rhodotorula spp. isolate 24–48 h in regular yeast agar plates (i.e., YM) at room temperature (20–25°C). (2) Transfer two full loopfulls of the biomass into a 1.5-ml tubes containing 500 μl of lysing buffer and the equivalent to 200 μl of 425–600 µm glass beads. (3) Vortex vigorously for 3 min, incubate at 65°C for 1 h and repeat vortexing (3 min). (4) Centrifuge suspensions for 15 min at 10000 rpm and preferably at low temperatures (5°C). (5) Collect supernatant and transfer to new tube; store at –20°C for up to 10–12 months. (6) For PCR studies dilute 1:600 in double distilled water in order to obtain a DNA solution ranging from 5 to 50 ng/ml; add 5 μl to the PCR mix and store at –20°C for up to 4–6 months. Alternatively, several commercial kits for yeasts genomic DNA extraction are available.

43.2.3  Detection Procedures 43.2.3.1 Identification and Genotyping of Rhodotorula spp. by Micro/ mini-satellite Primed PCR (MSP-PCR) The MSP-PCR typing method is a valuable tool for mycological laboratories due to its rapidity, high output and high strain discrimination potential. Suggested MSP-PCR primers for each medically relevant Rhodotorula species:

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Molecular Detection of Foodborne Pathogens

Table 43.2 List of Materials Needed for Different Molecular Procedures for the Detection of Rhodotorula spp Equipment

Culture incubator; Benchtop centrifuge; P1000 micropipette

Reagents

Labware and Consumables

A. DNA extraction from pure cultures of Rhodotorula spp. YM agar medium (3 g/l yeast extract, 5 g/l peptone, Disposable gloves (latex, vinile or nitrile); 10 g/l glucose, 3 malt extract, 20 g/l agar-agar); Eppendorf-like sterilized 1.5 ml micro tubes; Lysing buffer (50 mM Tris-HCl pH 8, 50 mM EDTA, Sterilized pipette tips for P1000 micropipette. 250 mM NaCl,and sodium dodecyl sulfate 425–600  µm washed (with diluted chloridric 1 % wt/vol); acid) glass beads

B. Typing of Rhodotorula spp. by MSP-PCR fingerprinting Disposable gloves (latex, vinile or nitrile); Thermal cycler; Taq DNA polymerase, MgCl2, 10 × PCR reaction Sterile pipette tips for P2/P10, P20/P200, PCR dedicated flow cabinet (optional); buffer, dNTPs, sterile water P1000 with filter (optional); Variable volume micropipettes (P2, P20, P200, MSP-PCR primer; 0.2 ml thin wall sterile tubes and 0.5 ml and P1000); Ethidium bromide or equivalent. sterile tubes. Horizontal electrophoresis system and power 1.4 % agarose gel supply. Molecular marker UV transiluminator and gel documentation system Loading dye C. Detection of Rhodotorula spp. employing species-specific PCR primers Thermal cycler; Disposable gloves (latex, vinile or nitrile); Taq DNA polymerase, MgCl2, 10 × PCR reaction PCR dedicated flow cabinet (optional); Sterile pipette tips for P2/P10, P20/P200, buffer, dNTPs, sterile water Variable volume micropipettes (P2, P20, P200, P1000 with filter (optional); Universal primer pairs and species-specific primer; and P1000); 0.2 ml thin wall sterile tubes and 0.5 ml Ethidium bromide or equivalent. Horizontal electrophoresis system and power sterile tubes. 1 % agarose gel supply. Molecular marker UV transiluminator and gel documentation system Loading dye D. Identification of Rhodotorula spp. by restriction of the amplified 5.8S-ITS region of the ribosomal rRNA operon Thermal cycler; Disposable gloves (latex, vinile or nitrile); Taq DNA polymerase, MgCl2, 10 × PCR reaction PCR dedicated flow cabinet (optional); Sterile pipette tips for P2/P10, P20/P200, buffer, dNTPs, sterile water Variable volume micropipettes (P2, P20, P200, P1000 with filter (optional); ITS1/ITS4 primers stock solutions; and P1000); 0.5 and 0.2 ml thin wall sterile tubes Endonucleases CfoI, HaeIII, and HinfI and respective Horizontal electrophoresis system and power with caps. buffers; supply. Agarose for routine use; UV transiluminator and gel documentation system Ethidium bromide or equivalent. 1 and 3 % agarose gels Molecular size marker (100–1000 bp) Loading dye

R. mucilaginosa: the core sequence of the phage M13 (5´-GAG GGT GGC GGT TCT-3´) or the synthetic oligonucleotide (GTG)5 provide enough species level resolution; R. minuta: same as R. mucilaginosa; R. glutinis complex: M13 primer allows detection of R. glutinis species complex, though discrimination of species within this group or strains may be achieved with the (GAC)5 primer. The MSP-PCR procedure is described below and materials are listed in Table 43.2. (1) Prepare PCR mixture containing 0.8 µM each primer, 2 mM of each deoxynucleotide, 3.5 mM MgCl2, 1 U of Taq DNA polymerase and 1 × PCR buffer, sterile double-distilled water up to the desired volume and 5 µl of the DNA sample (1:600 dilution) for a 25 µl reaction. (2) The cycling program consists of initial denaturation at 95°C for 5 min; 40 cycles of denaturing at 94°C for 45 sec, annealing at 50–55°C for 1 min and

extension at 72°C for 1 min; and a final extension at 72°C for 6 min. (3) Run gel electrophoresis on 1.4 % agarose gel, 0.5 × TBE buffer; at 90 V for 3.5 h. Include a molecular size marker on each gel, a mixture of DNA cleaved with HindIII and ΦX174 DNA cleaved with HaeIII is recommended. (4) Stain the gel in 0.5 µg/ml ethidium bromide aqueous solution. (5) Compare obtained MSP-PCR profiles with database or reference strains. 43.2.3.2 Detection of Rhodotorula spp. Employing Species-specific PCR Primers Species-specific oligonucleotide primers may be designed for PCR identification of known Rhodotorula species or group of species. The procedure uses standard PCR components and protocols (see Section 43.2.3.1) including DNA from the test

611

Rhodotorula

species and three primers: two universal external limiting primers (either universal D1/D2 region or ITS primers) and a species-specific internal primer. Species identification requires the amplification of a species-specific rDNA nucleotide segment that is significantly smaller than the non-target segment. Primers to be employed for the detection of R. mucilaginosa with species-specific PCR primers132: External universal primers: NL1 (forward): 5´-GCA TAT CAA TAA GCG GAG GAA AAG-3´ and NL4 (reverse): 5´-GGT CCG TGT TTC AAG ACG G-3´ Internal specific primer (forward) of the D1/D2 region: 5´-TCA GAC TTG CTT GCC GAG CAA TCG-3´.132 43.2.3.3 Identification of Rhodotorula spp. by Restriction of the Amplified 5.8S-ITS Region of the Ribosomal rRNA Operon (RFLP-ITS) A frequently used method for rapid identification of yeasts consists of PCR amplification of the ITS1-5.8S-ITS2 rDNA ITS, followed by digestion with restriction endonucleases and by separation by agarose gel elecrophoresis.33,140,141 Studies using the RFLP-ITS method for the identification of Rhodotorula yeasts are scarce, comprising only a few reports of wine166,167 and orange-juice spoilage yeasts.33 The RFLP-ITS profiles to date known corresponding to Rhodotorula spp. are the following: R. mucilaginosa strains typically show bands of the following molecular weights: ITS amplicon: ~600 bp, CfoI: 320 + 240 + 80, HaeIII: 425 + 215 and HinfI: 340 + 225 + 75 (Guillamon 1998, Sabate, 2002); whereas R. minuta has ITS amplicons of 660 bp and CfoI: 300 + 300, HaeIII: 400 + 215, HinfI: 345 + 215.166 R. glutinis complex, on the other hand, generates ITS amplicons of approximately 640–660 bp, and restriction fragments of variable sizes with all three enzymes (see database www.yeast-id. com). Unfortunately, some strains of R. glutinis complex share similar RFLP-ITS profiles with R. mucilaginosa. The only differences observed are seen in the fragments produced with HaeIII, which generally are 425 + 215 bp for R. mucilaginosa and 430 + 210 for R. glutinis complex.33,141 The protocol for the RFLP-ITS technique is described below and the required reagents, consumables, and equipment are listed in Table 43.2. The amplification/digestion procedure is applicable also directly from fresh yeast colonies for rapid identification (see below). (1) Prepare PCR mixture containing 0.5 µM each primer, 10 µM deoxynucleotides, 1.5 mM MgCl2 and 1 × PCR buffer, sterile double-distilled water up to the desired volume and 5 µl of the DNA sample (conveniently diluted) for a 25 µl reaction. For the alternative method of direct amplification from a single colony the cell material (picked with sterile tip or needle) is added to a reaction mixture of 25 µl (though mix components are at a concentration corresponding to 50 µl of final volume). After heating of the mixture containing the cell material for 15 min at 95°C, 1 U of Taq DNA polymerase is added in 25 µl double distilled water. (2) The cycling program consists of initial denaturation at 95°C for 5 min; 40 cycles of denaturing at 94°C





for 1 min, annealing at 55°C for 2 min and extension at 72°C for 2 min; and a final extension at 72°C for 10 min. (3) Digest the PCR products (15 µl or approximately 0.5–1.0 µg) without further purification with 1 U/µl of the restriction endonucleases CfoI, HaeIII, and HinfI at 37°C overnight. (4) Separate the PCR products and their restriction fragments on 1.4% and 3% agarose gels, respectively, in 0.5 × TBE buffer. (5) Stain the gels in 0.5 µg/ml ethidium bromide aqueous solution. (6) Estimate the sizes of PCR amplicons by comparison against a DNA length standard (e.g., 1000–100 bp ladder).

43.2.3.4 Identification of Rhodotorula spp. by rDNA Sequencing Analysis The rRNA genes are the preferred target for DNA sequencing. The main reasons are the availability of conserved and highly variable regions within the rRNA operon and its presence in multiple copies in fungal cells allowing high sensitivity, as well as accurate molecular identification for most yeasts.164,165 The 5´-end of the 26S rDNA D1/D2 domain (D1/D2) and the 5.8S-ITS region of the ribosomal rRNA gene (ITS) are the most frequently sequenced regions for yeast identification. Hence, large databases are available for nontaxonomist to quickly and accurately identify most known yeast species, as well as recognize new species, by sequencing approximately 600 nucleotides and doing a BLAST search in GenBank. In general, strains differing in no more than three nucleotide differences (0–0.5%) are considered conspecific, while strains showing six or more noncontiguous substitutions (1%) are separate species.168 Strains with intermediate nucleotide substitutions are also likely to be separate species, though further molecular and conventional characterization is needed for reassurance. On the other hand, the internal transcribed spacer regions ITS1 and ITS2, which are separated by the 5.8S gene of rDNA, are also highly substituted and often used for species identification, but for many species, ITS sequences provide no greater resolution than that obtained from 26S domains D1/D2.169,170 The required reagents, consumables, and equipment are listed in Table 43.2. (1) Prepare PCR mixture containing 0.5 µM each primer (see below), 10 µM deoxynucleotides, 1.5 mM MgCl2 and 1 × PCR buffer, sterile double-distilled water up to the desired volume and 5 µl of the DNA sample (diluted, sea above DNA extraction protocol) for a 25 µl reaction. For the alternative method of direct amplification from a single colony the cell material (picked with sterile tip or needle) is added to a reaction mixture of 25 µl (though mix components are at a concentration corresponding to 50 µl of final volume). After heating of the mixture containing the cell material for 15 min at 95°C, 1 U

612







Molecular Detection of Foodborne Pathogens

of Taq DNA polymerase is added in 25 µl double distilled water. (2) The cycling program consists of initial denaturation at 95°C for 5 min; 40 cycles of denaturing at 94°C for 1 min, annealing at 55°C for 2 min and extension at 72°C for 2 min; and a final extension at 72°C for 10 min. (3) The presence of PCR products must be confirmed by 1% gel electrophoresis. (4) Purification of the PCR product should be carried out using DNA purification kit following manufacturer´s indications. (5) Sequence PCR product using one or both of the primers either by sending samples to a DNA Sequencing Service or, if applicable, using your own Sequencer equipment following the manufacture´s protocol. (6) Visually correct sequences for nucleotide ambiguities or errors and compare to GenBank nucleotide database by using the BLASTN program (NCBI, Bethesda, MD).171

Type strains of medically relevant Rhodotorula species and their GenBank accession numbers for identification purposes are shown in Table 43.3. List of universal primers for rDNA sequencing: D1/D2 region: NL1 (forward): 5´- GCA TAT CAA TAA GCG GAG GAA AAG - 3´ and NL4 (reverse): 5´- GGT CCG TGT TTC AAG ACG G - 3´ primers.

ITS region: ITS1 (forward): (5’-TCC GTA GGT GAA CCT GCG G-3’) and ITS4 (reverse) (5’-TCC TCC GCT TAT TGA TAT GC-3’) primers. ITS1 region: ITS1 (forward): (5’-TCC GTA GGT GAA CCT GCG G-3’) and ITS2 (reverse) (5´-GCT GCG TTC TTC ATC GAT GC- 3´) primers. ITS2 region: ITS3 (forward) (5´-GCA TCG ATG AAG AAC GCA GC-3´) and ITS4 (reverse) (5’-TCC TCC GCT TAT TGA TAT GC-3’) primers.

43.3 Conclusions and Future Perspectives In spite of the large contact that humans have with Rhodotorula yeasts as a consequence of the broad distribution in natural and artificial environments of these microorganisms, a relatively low incidence of rhodotorulosis cases have been reported. This suggests that Rhodotorula infections are opportunistic and that the pathogenicity of these yeasts is limited by factors that are currently unknown and need investigation. Nevertheless, it is clear that Rhodotorula yeasts, in particular the species referred to in this chapter, must be considered medically relevant given that inmunocompromised patients are susceptible of developing fungeamia when exposed to such yeasts. The characteristic broad occurrence in natural and artificial environments of Rhodotorula species, especially of R. mucilaginosa, is most certainly a result of their physiological and metabolic plasticity. Such characteristic is also responsible for their frequent appearance in food and beverages. As

Table 43.3 The Type or Reference Strains for Medically Relevant Rhodotorula and other Pigmented Yeast Species Species

Type strain

D1/D2d

ITSd

R. mucilaginosa R. glutinis R. dairenensis Rh. sphaerocarpum Rh. toruloides Rh. kratochvilovae Rh. diobovatum Rh. babjevae R. graminis R. minuta Other Sporidiobolus johnsonii a Sporidiobolus salmonicolor b Sporidiobolus metaroseus c Sporidiobolus pararoseus

CBS 316T CBS 20T CBS 4406T CBS 5939T CBS 14* CBS 7436T CBS 6085T CBS7808T CBS 2826T CBS319T

AF070432 AF070430 AF070429 AF070432 AF207884 AF071436 AF070421 AF070420 AF070431 AF189945

AF444541 AF444539, AF387775 AF444501 AF444499 AB049028 AF444520 AF387782, AF444502 AF444542 AF444505 AF190011

CBS 5470T CBS 490T CBS 7687T CBS 491T

AF070435 AF070439 EU003461 AF189977

Notes: R.: Rhodotorula, Rh.: Rhodosporidium.   Formerly: Sporobolomyces holsaticus. b  Formerly Sporobolomyces salmonicolor. c  Formerly: Sporobolomyces roseus. d  GenBank accession number. * Not the type strain but a widely used reference strain. a

EF592109 AF387784 EU003482 AY015429

613

Rhodotorula

already discussed, food, and beverages can be a significant source of Rhodotorula yeasts thus posing an additional and probably underestimated risk for susceptible patients. Future studies should aim to the clarification of the contribution of foods and beverages to opportunistic yeast infections by Rhodotorula spp. In the mean time, special attention should be paid to food and beverages given to patients with any type of immune-compromising condition. In recent years, several yeast species (mainly of the genera Rhodotorula and Cryptococcus) have been claimed to be useful for the biological control of postharvest diseases in storage fruits and vegetables due to their antagonistic activities against common plant pathogens.172 Among the Rhodotorula species reported as potential biocontrol agents R. mucilaginosa,172 R. glutinis 173–177 and R. minuta178 are included and have been tested with promising results. Concern arises regarding the use of these Rhodotorula species in food for human consumption, given that an increase in their natural loads of yeasts that can act as opportunistic pathogens might be expected. Further studies devoted to assess the viability of such yeasts when treated products reach the consumers should be carried out. Future investigations must also focus on the factors that favor infection by Rhodotorula yeasts, and on the search for phenotypic and genotypic characteristics that may help discriminate potential pathogenic strains from those that are not. In this task, molecular genotyping tools will play a significant role by allowing the rapid screening of clinical and environmental isolates for the detection of molecular traits proper of more virulent strains. The increasing importance of Rhodotorula yeasts as infective agents makes of the possibility to distinguish between potentially pathogenic and harmless strains on the basis of genetic traits a pressing need. Molecular techniques for the detection, identification, and typing of potentially pathogenic strains of Rhodotorula must provide rapid and accurate identification and discrimination from taxonomically related species. For the case of Rhodotorula, a requisite for such molecular technique is that it must grant precise species differentiation within the R. glutinis complex. This will be the only way to determine which of these species can actually cause disease and in the hypothetical case that all did, which has the higher incidence of infections. Such epidemiological investigations are needed and are important for a correct circumscription of the Rhodotorula species that can behave as pathogens, thus allowing the assessment of their antifungal susceptibilities. The implementation of molecular biology techniques in the clinical laboratory for the routine detection of Rhodotorula spp. is vital for a prompt and precise diagnosis, and the subsequent application of the appropriate antifungal treatment. Current conventional methodologies based on phenotypic criteria do not provide the required specificity, often leading to inconclusive identifications or even to incorrect ones. The techniques proposed in the present chapter have been used extensively in the identification and typing of Rhodotorula strains from different sources including clinical, and may be perfectly implemented for identification

and/or epidemiological purposes in the clinical laboratory. As discussed above, rDNA sequence analysis represents a powerful molecular tool for the rapid and accurate diagnosis of Rhodotorula spp. as well as of many other infections caused by fungi, and is probably the best choice for routine clinical practice. However, each clinical laboratory will determine which molecular technique can be used on the basis of its workflow, workload, and available financial resources. An additional advantage of molecular techniques over conventional ones is that the direct detection of pathogens from clinical specimens becomes possible. Such technology has been already investigated for several pathogenic yeasts, though mostly of the genera Candida 135,179–181 and Cyrptococus153,182 from various types of specimens. The molecular techniques that have been to date evaluated for culture-independent diagnosis of yeast infections were based on real-time PCR,179,135,181 DNA microarrays,156,180 FISH,182 and Luminex®153 technologies. However, to the best of our knowledge, none of these techniques have been yet tested for detection of Rhodotorula spp. directly from human specimens. Thus, future investigations and developments in this area should take account of Rhodotorula species due to their increasing importance as opportunistic mycoses-producing yeasts.

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618 177. Xu, X., Qin, G. and Tian, S. Effect of microbial biocontrol agents on alleviating oxidative damage of peach fruit subjected to fungal pathogen. Int. J. Food Microbiol., 126, 153. 2008. 178. Patiño-Vera, M. et al. Pilot-scale production and liquid formulation of Rhodotorula minuta, a potential biocontrol agent of mango anthracnose. J. Appl. Microbiol.,99, 540, 2005. 179. Salvarangan, R. et al. Rapid identification of commonly encountered Candida species directly from blood culture bottles. J. Clin. Microbiol., 41, 5660, 2003. 180. Spiess, B. et al. DNA Microarray-based detection and identification of fungal pathogens in clinical samples from neutropenic patients. J. Clin. Microbiol., 45, 3743, 2007. 181. Vollmer, T. et al. Evaluation of novel broad-range real-time PCR assay for rapid detection of human pathogenic fungi in various clinical specimens. J. Clin. Microbiol., 46, 1919, 2008. 182. Wilson, D.A. et al. Multicenter evaluation of a Candida albicans peptide nucleic acid fluorescent in situ hybridization probe for characterization of yeast isolates from blood cultures. J. Clin. Microbiol., 43, 2909, 2005. 183. Herzberg, M., Fischer, R. and Titze, A. Conflicting results obtained by RAPD-PCR and large-subunit rDNA sequences in determining and comparing yeast strains isolated from flowers: a comparison of two methods. Int. J. Syst. Evol. Microbiol., 52, 1423, 2002. 184. Vermitsky, J.-P. et al. Survey of vaginal-flora Candida species isolates from women of different age groups by use of

Molecular Detection of Foodborne Pathogens species-specific PCR detection. J. Clin. Microbiol., 46, 1501, 2008. 185. Walsh, T.J. et al. PCR and single-strand conformational polymorphism for recognition of medically important opportunistic fungi. J. Clin. Microbiol., 33, 3216, 1995. 186. Sugita, T. et al. Sequence analysis of the ribosomal DNA intergenic spacer 1 regions of Trichosporon species. J. Clin. Microbiol., 40, 1826, 2002. 187. Sugita, T. et al. Sequence diversity of the intergenic spacer region of the rRNA gene of Malassezia globosa colonizing the skin of patients with atopic dermatitis and healthy individuals. J. Clin. Microbiol., 41, 3022, 2003. 188. Biswas, S.K. et al. Molecular phylogenetics of the genus Rhodotorula and related basidiomycetous yeasts inferred from the mitochondrial cytochrome b gene. Int. J. Syst. Evol. Microbiol., 51, 1191, 2001. 189. Trnovsky, J. et al. Rapid and accurate identification of Candida albicans isolates by use of PNA FISHFlow. J. Clin. Microbiol., 46, 1537, 2008. 190. Montero, C.I. et al. Evaluation of Pyrosequencing® technology for the identification of clinically relevant non-dematiaceous yeasts and related species. Eur. J. Clin. Microbiol. Infect. Dis., DOI: 10.1007/s10096-008-0510-x, 2008. 191. Diaz, M.R. et al. Molecular sequence analyses of the intergenic spacer (IGS) associated with rDNA of the two varieties of the pathogenic yeast. Cryptococcus neoformans, Syst. Appl. Microbiol., 23, 535, 2000.

44 Saccharomyces

Franca Rossi and Sandra Torriani Università degli Studi di Verona

Contents 44.1 Introduction.....................................................................................................................................................................619 44.1.1 General Characteristics......................................................................................................................................619 44.1.2 S. cerevisiae variant S. boulardii...................................................................................................................... 621 44.1.3 S. cerevisiae as an Emerging Pathogen............................................................................................................ 622 44.1.4 Virulence Attributes of Pathogenic Strains...................................................................................................... 624 44.1.5 Molecular Identification and Characterization of S. cerevisiae Strains........................................................... 625 44.2 Methods.......................................................................................................................................................................... 627 44.2.1 Reagents and Equipment ................................................................................................................................. 627 44.2.2 Sample Treatment............................................................................................................................................. 627 44.2.3 Detection Procedures . ..................................................................................................................................... 631 44.2.3.1 Detection of S. cerevisiae by Real-time PCR.................................................................................. 631 44.2.3.2 Identification of S. cerevisiae by Restriction of the Amplified 5.8S-ITS Region of the Ribosomal rRNA Operon...................................................................................................... 632 44.2.3.3 Typing of S. cerevisiae by Microsatellite Amplification................................................................. 632 44.3 Conclusions and Future Perspectives............................................................................................................................. 633 References.................................................................................................................................................................................. 633

44.1  Introduction 44.1.1 General Characteristics The genus Saccharomyces (from Latin meaning sugar fungi) consists of multiple species of yeasts, which are noted for the roles they play in food production. The budding yeast Saccharomyces cerevisiae is included in the Saccharomyces sensu stricto yeast clade with the closely related species Saccharomyces bayanus, Saccharomyces cariocanus, Saccharomyces kudriavzevii, Saccharomyces mikatae, Saccharomyces paradoxus, and S. pastorianus (which is considered a hybrid of S. cerevisiae and S. bayanus). This group of species belongs to the fungal phylum Ascomycota, subphylum Saccharomycotina. The latter is composed of three clades among which the “Saccharomyces complex,” covering the genera Saccharomyces and Kluyveromyces, is the most extensively studied. The phylogeny of this group of species was defined by multigene sequence comparison1 and by comparative hybridization of genomic DNA on S. cerevisiae cDNA microarrays.2 Based on the complete genome sequences, S. bayanus, S. mikatae, S. kudriavzevii and S. paradoxus are very close relatives of S. cerevisiae.3 The species S. cerevisiae, S. bayanus, S. mikatae and S. paradoxus are not prezygotically isolated: F1 hybrids can be produced through mating of haploid cells.4–6 These species are, however, postzygotically isolated because the F1 diploid cells are mostly sterile, producing 1% or fewer of viable spores. This is partly due to the mismatch repair system that interferes with proper recombination during meiosis in the F1 hybrids of

S. cerevisiae and S. paradoxus.4 A common ancestor of the subgroup “Saccharomyces complex” comprising S. cerevisiae underwent whole genome duplication.7 This event was followed by the loss in one member of most of the duplicated gene pairs and concomitant speciation. Ribosomal protein genes and regulatory genes are those preferentially retained in duplicate. Some of the duplicate genes were maintained in consequence of neofunctionalization; among these, genes involved in adaptation to low oxygen environments are found. In some species, and also in S. cerevisiae, this resulted in the abilities to consume glucose preferentially via fermentation rather than respiration, to grow in total absence of oxygen and to be induced to form respiratory deficient mitochondrial mutants (“petites”).8 Other genes retained in duplicate in S. cerevisiae are involved in carbohydrate metabolism and are differentially expressed in response to oxygen or glucose availability.9 Further adaptations to the “fermentative lifestyle” occurred specifically in an ancestor of the Saccharomyces sensu stricto species, with duplications of ADH and PDC genes.10 Although fermentation is widely regarded as a losing strategy owing to the relatively low output of ATP, in S. cerevisiae it is such a rapid process that optimization of glycolysis allows to synthesize a similar number of ATP molecules per second as in the aerobic metabolism.11 The ability to produce ethanol during fermentation is not unique to the Saccharomyces species. The uniqueness of Saccharomyces rather resides in its ability to produce and to tolerate high levels of ethanol, which may later be utilized as a source of energy once that glucose is depleted.12 Not surprisingly, 619

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S. cerevisiae is used worldwide in the food industry as principal agent of fermentation in naturally leavened bakery products, alcoholic beverages like wines, beers, disparate fermented juices, and distillates. It is also consumed as a nutritional supplement for its high concentration of the B vitamins, minerals, and cofactors. This yeast species can be found in nature on various plants and in the soil.13–15 In studies aimed at defining the source from which it contaminates grapes in the vineyards and the winery environment it has been isolated from single rotting fruits, but only from 0.1% of berries with undamaged surface. S. cerevisiae was also isolated from the bodies of insects, most frequently from the honey bees, but also from wasps, and Drosophila. These are supposed to be the carriers that transfer the yeast to the interior of damaged fruits when descending for nutrition, since it has been demonstrated that S. cerevisiae is not transported by air in the field environment. Indeed, the only ascertained case in which Saccharomyces cells are transmitted through air13 when it is spread from the foam of fermenting must.16 During wine fermentation the species S. cerevisiae complete the process initiated by the less ethanol-tolerant genera Kloeckera, Hanseniaspora, Candida, and Pichia. The high level of ethanol produced, along with the anaerobic conditions, low pH, and the osmotic stress, eliminates other microorganisms. As a result, strains of S. cerevisiae, indigenous or added as starter, are essentially the only organisms remaining alive at the end of a fermentation.17 Nucleotide variation in 81 isolates originating from natural and artificial fermentations, tree exudates, and immunocompromised patients from Europe, Africa, Southeast Asia, and North America supports the hypothesis that “domesticated” strains derive from natural populations rather than the other way around.15 In this phylogenetic study based on diversity at five loci in a sequence of 7 kb, half coding and half noncoding, 184 polymorphic sites were identified. The genealogical relationships among these strains did not offer a clear distinction between domesticated and nondomesticated. The phylogenetic tree rather suggested that lineages at the root were derived from tree exudates in North America and Africa. The comparison of oak-associated and vineyardassociated yeasts suggested that habitats and geographical distance may play a role in shaping the patterns of genetic diversity as they cluster in different phylogenetic groups.15 Life cycle, cell structures assembly, metabolic pathways, gene function, and expression regulation of S. cerevisiae are the most well known among eukaryotes, since this organism constituted for decades the model system for studying these aspects thanks to the relatively small and compact genome, comprising 16 chromosomes of sizes 200–2,200 kb in the haploid stage, and ease of genetic transformation and manipulation. Knowledge about the physiology of this yeast has increased after strain S. cerevisiae S288c genome sequencing and annotation, that unravelled the presence of about 6,000 genes covering 72% of the total sequence, of which 3.8% are introns, was completed.18 Chromosomes contain movable DNA elements, retrotransposons, that vary in number and position in different strains of S. cerevisiae, with

Molecular Detection of Foodborne Pathogens

most laboratory strains having approximately a total of 30 copies subdivided in the retrotransposon types Ty1–Ty5. These are 5.3–5.7 kb in size and are constituted by retroviral DNA flanked by ~350 bp terminal repeats (LTRs). Retrotransposons Ty1 and Ty2 are flanked by δ sequences. Other nucleic acid types are mitochondrial DNA, that encodes components of the mitochondrial translational machinery and approximately 15 % of the mitochondrial proteins, and the 2-µm circle plasmids, present in most strains of S. cerevisiae, that apparently function solely for their own replication. Almost all S. cerevisiae strains contain dsRNA viruses, that constitute approximately 0.1% of total nucleic acid. RNA viruses include three families with dsRNA genomes, L-A, L-BC, and M. Two other families of dsRNA, T, and W, replicate in yeast but so far have not been shown to be viral. M dsRNA encodes a killer toxin, and L-A encodes the major coat protein and components required for the viral replication and maintenance of M. The two dsRNA, M, and L-A, packaged separately with the common capsid protein encoded by L-A, are transmitted cytoplasmically as virus-like particles during vegetative growth and conjugation. KIL-o mutants, lacking M dsRNA and consequently the killer toxin, are readily induced by growth at elevated temperatures, and by chemical and physical agents. L-B and L-C (collectively denoted L-BC), similar to L-A, have a RNA-dependent RNA polymerase and are present in intracellular particles. S. cerevisiae also contains a 20S circular single-stranded RNA that appears to encode an RNA-dependent RNA polymerase, that acts as an independent replicon and is inherited as a nonMendelian genetic element. The life cycle of S. cerevisiae alternates between diplophase and haplophase. Both ploidies can stably propagate asexually. The sizes of haploid and diploid cells vary with the phase of growth and from strain to strain. Vegetative cell division of yeast characteristically occurs by axial or bipolar budding, in which a daughter cell is initiated as an out growth from the mother cell (phase S), followed by nuclear migration and division (phases G2-and M), cell-wall formation and cell separation (phase G1). In the budding region a scar is left on the mother cell surface. In heterothallic strains, haploid cells are of two mating types, a and α. Mating of a and α cells results in a/α diploids that are unable to mate but can undergo meiosis upon starvation. The spores of about 70% of natural isolates are able to switch mating type; these are referred to as homothallic strains.19 The wild-type allele of the gene HO for homothallism encodes a site-specific endonuclease that promotes mating type switching from a to α or vice versa. HO is transcribed only in the mother cell, so that when a homothallic spore divides, the mother cell switches mating type and immediately fuses with the newly produced daughter cell.20 The four haploid products resulting from meiosis of a diploid cell are contained within the wall of the mother cell (the ascus). Asci have higher resistance than vegetative cells to environmental stress, such has heat and dehydration. Digestion of the ascus and separation of the spores by micromanipulation yield the four haploid meiotic products that can be analyzed singularly for tracing gene segregation and linkage.

621

Saccharomyces

The fact that about 27% genes are expressed in a way correlated with growth rate, independent of the identity of limiting nutrient, i.e., glucose, nitrogen, sulphur, or phosphate, emerged from a study based on DNA microarray analysis of cultures grown with different nutrient limitations. A simple linear relationship was found, between steady-state growth rate and the fraction of unbudded cells. A positive correlation exists between growth rate and the transcription of genes of both cytosolic and mitochondrial translation machinery. Genes related to functions associated with oxidative energy metabolism, especially those carried out in peroxisomes, namely, fatty acid metabolism, oxygen metabolism and autophagy, are negatively correlated with growth rate. The hypothesis to explain this is that it serves to protect the cell from the longer exposure to oxidative metabolism necessary to produce enough energy at lower growth rates.21 Array-based comparative genomic hybridization highlighted an abundant genetic variation among the 14 S. cerevisiae wine and laboratory strains assayed both at the level of nucleotide polymorphism and gene copy number.22 Preferential location of gene deletions were the subtelomeric regions, where many genes involved in C-compound metabolism, transport, and fermentation are found. Indeed, these are also rich in redundant sequences potentially involved in nonhomologous recombination to generate differences in gene copy number.23 S. cerevisiae also displays a variety of colony morphologies. While it is mainly known in the laboratory as a unicellular species with ellipsoid cells that divide by multilateral budding and form smooth colonies on solid medium, wine yeasts can also form more complex colonies by modifying their budding patterns and forming multicellular pseudohyphae,24 a process also called “pseudofilamentation.” Other phenotypes include the formation of biofilms25 and complex “fluffy”-structured colonies with cells connected by extracellular glycosylated matrix material.26 These alternative morphologies are hypothesized to be adaptive responses to environmental signals. For instance, the formation of pseudohyphae is induced by nitrogen deprivation and allows the cell to penetrate in the solid medium for nutrients.24 The formation of “fluffy” colonies might also represent an adaptive response against desiccation by forming small channels for nutrient and water flow.27 Filamentous growth is also of medical interest because, in cases of human infection, this mode of propagation is thought to allow the penetration of physiological tissues.28 A screen of 1,026 strains of yeast isolated from Italian vineyards showed that roughly half of the strains were able to show invasive filamentous growth when starved for nitrogen. Also, 2.5% of the strains had a filigreed (“rough”) or fluffy colony phenotype. Furthermore, several strains also showed differences in the treatments that induce modification of growth morphology, indicating the presence of genetic variation in the response to stimuli that can produce filamentous growth.29 In a study on the segregants from a Montalcino strain M28 it was observed that filigreed progeny expressed MEP1, MEP2, and MEP3 ammonia transporters at higher levels, while smooth colony progeny exhibited higher expression of amino acid transporters.30

44.1.2  S. cerevisiae variant S. boulardii The S. cerevisiae variant “S. boulardii” (patent strain held in the American Type Culture Collection, ATCC), first isolated from litchi fruit in Indochina, once considered a separate species and later reclassified as S. cerevisiae Hansen CBS 5926,31,32 is used worldwide as a biotherapeutic agent. “S. boulardii” has been used in Europe as probiotic preparations under the commercial names Ultra-Levure (Biocodex, France), Precosa® (Logic aps., Denmark), Codex (Zambon Farmaceutici, Italy), and has been proposed for use also in the United States, where it has been studied for oral administration in lyophilized form the by Biocodex Inc. (Seattle, WA). “S. boulardii” is also utilized as feed additive, an example is Levucell SB® (Euro Tier, Germany) and as food supplement. The correct molecular identification of “S. boulardii” as S. cerevisiae was obtained by the microsatellite amplification of Hennequin et al.:33 positive PCR results obtained with “S. boulardii” isolates strongly suggested that “S. boulardii” is a strain of S. cerevisiae rather than a different species. Before the assessment of this technique it was not possible to identify “S. boulardii” in a routine manner. Phenotypic traits such as the lack of galactose assimilation and the lack of α-glucosidase activity are characteristics of “S. boulardii”. However, none of these traits allows a definitive identification, and confirmation requires multiple enzymatic restrictions of mtDNA. Interestingly, even the highly discriminatory randomly amplified polymorphic DNA (RAPD) method failed to differentiate between isolates of S. cerevisiae and “S. boulardii”.34 In contrast, it was found that microsatellite repeat sequence (CAG)9 in one of the loci tested is specific for this strain and thus constitutes a rapid alternative for an accurate identification of “S. boulardii”. Moreover, this strain can be differentiated from S. cerevisiae reference and clinical strains by microsatellite polymorphism in other five genetic loci, according to Malgoire et al.35 “S. boulardii” is highly effective either as a preventative agent or for the treatment of several types of acute diarrhea and colitis, like antibiotic-associated diarrhea, diarrhea in adults and children infected with Clostridium difficile, for diarrhea in human immunodeficiency virus-infected patients, traveller’s diarrhea, diarrhea in tube-fed patients, relapses in Crohn’s disease.36–39 The protective effects against C. difficile, entheropathogenic Escherichia coli and Vibrio cholerae are partly explainable with the inactivation of toxins by proteases, aminopeptidases, protein phosphatases secreted by the yeast in the intestinal lumen. Lyophilized preparations of “S. boulardii” also exert trophic effects on the small intestine mucosa that are mediated by the release of the polyamines spermidine and spermine in the endoluminal compartment. Moreover, “S. boulardii” improves the absorption of glucose by secretion of high amounts of sucrases and by the stimulation of disaccharidases and of glucose and electrolyte uptake by the sodium glucose cotransporter (SGPT) in the enterocytes. In addition, “S. boulardii” has been demonstrated to enhance the specific IgG and IgA antibody response to C. difficile toxin A and to Salmonella and Yersinia antigens. Since IgA

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production is the main defence of the intestinal barrier against invasion of the body by pathogens and toxins, this effect explains in part the protective effects of “S. boulardii.”40–42 Moreover, the inflammatory bowel disease symptoms are attenuated by “S. boulardii,” that is capable of inducing the migration and accumulation of T-cells, responsible of amplification of the inflammation reaction, in the mesenteric lymph nodes instead than in the inflamed colon.43 Positive immunologic effects exhibited also by other S. cerevisiae strains are the reduction of proinflammatory cytokine IL-1α expression in IPEC-J2 cells and diminution of IL-8 expression mediated by NF-Kβ in ΗΤ-29 colonocytes.44,45

44.1.3  S. cerevisiae as an Emerging Pathogen Though S. cerevisiae is commonly perceived as a harmless microorganism given its 1,000-year association with food and beverage production, numerous case reports of human infections caused by this yeast exist in the medical literature. These occurred with a broad range of clinical manifestations, from fungemia, pneumonia, and deeply invasive dissemination to localized infections in specific organs. Virulent S. cerevisiae possess the capability to colonize and persist in various normally sterile body sites (e.g., the lung and blood) and, consequently the ability to survive and disseminate in vivo. Local genitourinary infections may occur as a consequence of hormonal or mechanical factors in mildly immunocompromised or immunocompetent persons, while serious gastrointestinal or systemic infections have been observed in immunosuppressed patients. Since this microorganism is a common saprophyte, biopsy, and pathologic confirmation of infection are often necessary for a definitive diagnosis. Regarding the therapy of the infections caused by this yeast, it must be taken into account that S. cerevisiae exhibits low susceptibility to amphotericin B and to azole derivatives. Although breakpoints have not been defined for S. cerevisiae, MIC90 for fluconazole and traconazole are considered to be within the susceptible dose dependent range defined for C. albicans. Although data remain scarce, voriconazole seems to exhibit good efficacy against S. cerevisiae, with a MIC90 of 0.25 mg/l. A recent publication reports the successful outcome of voriconazole therapy for a patient with “S. boulardii” who did not respond to initial treatment with fluconazole.46 S. cerevisiae infections are apparently rare, however this yeast is currently considered an emerging pathogen in part as an effect of the refinement of yeast species identification methods but also for new patient predisposing factors. Indeed, though S. saccharomyces infections were observed in immunocompetent patients, the increase of incidence is a consequence of the increase of the population immunocompromised for underlying chronic or debilitating diseases, increased use of immunosuppressive drugs or broad spectrum antibiotics, as well as parenteral nutrition, use of intravascular catheters and, finally, the increasing numbers of HIV-infected individuals. Infections with lethal outcome were observed in patients with critical conditions such as

Molecular Detection of Foodborne Pathogens

AIDS, neoplastic, and metabolic diseases, intestinal ­surgery. Many cases were related to “S. boulardii” therapy. As a result, European Food Safety Authority (EFSA) recommended that “S. boulardii” should certainly be controindicated for patients of fragile health, as well as for patients with a central venous catheter in place.47 On the basis of data revealed in the literature survey presented by Enache-Angoulvant and Hennequin,48 invasive Saccharomyces infections remain rare, although the incidence has significantly increased since the 1990s. Risk factors associated with invasive S. cerevisiae infections are similar to those reported elsewhere for invasive candidiasis, except for treatment with a probiotic containing “S. boulardii.” It is important to emphasize the role of this biotherapeutic agent, because it was responsible for 40.2% of invasive S. cerevisiae infections reported in the literature. Although S. cerevisiae has been isolated from the digestive tract, vagina, skin, and oropharynx of healthy hosts, to date it has not been clearly demonstrated whether this is a true commensal species of human intestinal flora or if its colonization is transient and related to food consumption. Indeed, persistent digestive implantation in healthy volunteers has never been described. Thus, the portal of entry for invasive infections is mainly supposed to be digestive. One S. cerevisiae infection case in an immunocompetent patient, whose unique predisposing factor was the ingestion of health food containing viable yeasts, supports this hypothesis. The patient developed recurrent fever, malaise with nausea, and night sweats. Yeasts were isolated from bone marrow and urine specimens. The patient recovered when he stopped ingesting health food containing yeasts.48 In some reported cases, the S. cerevisiae strains responsible were winery resident strains or bakery strains.49–51 De Llanos et al.52,53 described the first isolation of a bakery strain from blood. Saccharomyces infection is clinically indistinguishable from invasive candidiasis, notably because chorioretinitis (fluffy yellow exudates) and oesophagitis (yellow-white plaques on an erythematous background) are present in both conditions, and is even indistinguishable from infections due to other microorganisms, such as endocarditis. In addition, patients have nonspecific symptoms that could be related to the underlying disease, and the diagnosis of Saccharomyces infection is often unexpected. Fever, when noted, is present in 75.2% of patients. The occurrence of deep-site involvement with histological documentation also underlines the pathogenicity of such yeasts. In these cases, yeasts were most often associated with necrosis and granulomatous reaction. For the establishment of the epidemiological trend, virulence levels and mechanisms, 92 documented cases of proven invasive S. cerevisiae infection were taken into account.48 Only proven infection, based on the definitions of the Invasive Fungal Infection Cooperative Group of the European Organization for Research and Treatment of Cancer-Mycosis Study Group of the National Institute of Allergy and Infectious Diseases cooperative group: fungemia, isolation from normally sterile fluids (e.g., pleural and synovial fluids), and deep-site infections either histologically proven for sites susceptible to

Saccharomyces

colonization (lungs, peritoneum, oesophagus), were considered. Fifteen cases (16.3%) were diagnosed before 1990, and 76 cases (82.6%) were diagnosed after 1990. All patients had at least one condition facilitating the development of invasive fungal infection. Intravenous catheter use (by 47 patients) and previous receipt of antibiotic therapy (by 45 patients) were the most frequently reported predisposing factors. “S. boulardii” was considered to be the etiologic agent in 37 cases. Molecular identification was performed in 23 cases, mainly by means of restriction-enzyme analysis of mtDNA. Among the 37 patients with presumptive “S. boulardii” infection, five did not take a probiotic containing “S. boulardii” at the time of diagnosis. However, the similarity of the genotypic profile between “S. boulardii” isolates from patients and from probiotic preparations strongly suggested nosocomial acquisition, with catheters being a likely portal of entry because of possible contamination through hand transmission. The same explanation is given for cases of S. cerevisiae infection in haematological patients because they are supplied exclusively with sterile food, without raw vegetables. Moreover, identical isolates from patients concurrently hospitalized in the same unit have been described. Compared with patients infected with S. cerevisiae, patients infected with “S. boulardii” were more likely to have digestive tract disease (5.6% vs. 58.3%), to have an intravenous catheter (29% vs. 83.8%), and to be hospitalized in an intensive care unit (0.05% vs. 32.4%). In contrast, the frequency of immunocompromise was lower among patients infected with “S. boulardii” than among patients infected with S. cerevisiae (25% vs. 58.5%). Saccharomyces organisms were most frequently isolated from blood (72 patients), either exclusively (63 patients) or with involvement of other organ(s) (nine patients). “S. boulardii” was exclusively isolated from blood. Concomitant isolation of S. cerevisiae and a second microorganism, mainly enterobacteria, was reported in seven cases of fungemia. Patient 10 received a post-mortem diagnosis of disseminated infection, although positive blood culture results were not recorded before death. In the remaining 17 cases, the infection was localized to a single organ. Overall, the main sites were the lungs (for eight patients) and the heart valves (for six patients). Patients with “S. boulardii” infection had a better prognosis than patients with S. cerevisiae infection (73% vs. 55.5%). The rates of favorable outcome among immunocompromised patients (65.5%) and immunocompetent patients (73.8%) did not differ significantly. There was no significant difference in the rates of favorable outcome between patients treated with intravenous amphotericin B (77.7%) and patients treated with fluconazole (60%). Favorable outcome may also be related to the low virulence of this species, a hypothesis that is supported by the absence of significant difference in outcome between immunocompromised and immunocompetent patients. The several documented reports of systemic infections like fungemia and septicemia with “S. boulardii” as etiological agent resulted from oral administration both in immunocompromised and immunocompetent patients. Data of microsatellite typing allowed to confirm that Saccharomyces

623

fungemias diagnosed in patients treated with “S. boulardii” therapy (eight cases) were indeed due to this particular strain.33 Though the level of virulence of “S. boulardii” is modest, it raises the question of the wisdom of using a potential pathogen as a therapeutic agent, especially in immunocompromised patients, in light of the demonstration that uptake of this organism from the gut lumen and translocation across the mucosa may occur.54 Moreover, administration of this probiotic is a risk for patients in the same care units as those that receive the treatment, indeed cases of fungemia concurrent to the administration of “S. boulardii” were registered and attributed to the hospital environment contamination.55 De Llanos et al.52 also found that among four isolates from patients with S. cerevisiae systemic infection one HinfI mtDNA restriction pattern and δ sequences amplification profile combination coincided with “S. boulardii” and could derive from use of this probiotic in a paediatric intensive care unit. In addition, it has been reported that the degree of virulence of “S. boulardii” isolates can vary and is therefore not predictable. Indeed, two isolates from different “S. boulardii” batches exhibited an intermediate degree of virulence compared with virulent and avirulent isolates of S. cerevisiae but also significantly differed in virulence in the same direction in two different models. This suggests that different batches of the preparation may not be uniform and may differ in a gene or genes that determine virulence. Strain variation is also indicated by the fact that the initial publication described a sporulating yeast, whereas the isolates examined more recently were asporogenous, as shown by repeated testing.31 Overall the epidemiology of human Saccharomyces infections remains still poorly understood although methods relying on genetic molecular markers have been assessed to distinguish strains. A study regarding DNA typing of 60 clinical and nonclinical isolates of S. cerevisiae reported that EcoRI DNA digestion profiles of clinical isolates were very heterogeneous, exhibiting little clonality. However, it was possible to subgroup these isolates into two clusters, A, and B, on the basis of the presence or absence of a 3.0-kb band on agarose gel electrophoresis of EcoRI-digested DNA and a statistical association of virulence with the group A DNA type was found.56 The comprehensive review of published reports of invasive Saccharomyces infections performed by Enache-Angoulvant and Hennequin48 had also the scope of generating hypotheses for predisposing preventive strategies. Epidemiologic and clinical differences characteristics between S. cerevisiae and “S. boulardii” invasive infections were revealed. Significant particularities of “S. boulardii” infections include higher frequency of intensive care unit hospitalization, presence of an indwelling catheter, and intestinal disease. This may either correspond to specific predisposing factors or, more likely, may reflect conditions associated with treatment with a probiotic containing “S. boulardii,” such as severe digestive disease. The lack of organ involvement and the better prognosis could be due to decreased “S. boulardii” virulence. Another explanation could be the ability of patients at risk

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to partly control the infection, given the lower frequency of immunocompromise in this group. Isolates of S. cerevisiae display a continuum of virulence potential and virulence in mice, defined as the capacity for the organism to proliferate in the brain and/or persist for an extended period of time,57 significantly associated with clinical strains.31 De Llanos et al.52 found that the δ sequences amplification profile combination of a S. cerevisiae isolate from an infection case was common to two commercial bread-making strains. The two molecular types were also found similar to several clinical isolates from feces, pharynx, and vagina previously characterized.58 These results importantly underlined that some S. cerevisiae food strains can reach the blood stream and create concern, considering the high number of people frequently exposed to bread-making strains. S. cerevisiae is responsible for symptoms mimicking Candida vaginitis.59 In an investigation carried out in an Italian hospital S. cerevisiae was implicated in 0.4% of cases (19 of 4,943) occurring during a 16 month experimental period.60 Medical literature reports describe cases of vaginal infections directly caused by S. cerevisiae strains used in domestic or professional bread-making.50,51 Vaginal isolates were associated to the group B endonuclease digestion DNA profile type of Clemons et al.56 with significant probability.60 The approach to successful management of S. cerevisiae infections is necessarily individual, but includes antifungal chemotherapy and, intuitively, the removal of potentially colonized intravascular catheters. Other adverse effects of S. cerevisiae on consumer’s health are allergic reactions mostly consisting of atopic dermatitis to yeast cell components. However, IgE immunoblotting demonstrated that S. cerevisiae antigens are not present in wine, beer, and bakery.61

44.1.4  Virulence Attributes of Pathogenic Strains Some fungal properties are frequently associated with pathogenesis, for example, the ability to grow at high temperatures (38–42°C), adherence and the ability to invade host cells, morphological versatility, including switching, and the ability to produce and secrete degradative enzymes (proteinase and phospholipase), which could play a role in blocking capillaries, a fungal characteristic associated with animal mortality.62 A key property of Saccharomyces strains for being potentially pathogenic is the maximum growth temperature: the ability to grow at 37°C showed no significant association with virulence, however, growth at 39°C was significantly associated with virulence. The ability to grow at 42°C was found almost exclusively in clinical strains (ten of 15 clinical strains were able to grow at 42°C, vs. one of ten nonclinical strains) and was tightly associated with virulence.63 This may suggest that any thermotolerant (viability at 37°C) S. cerevisiae strain is able to transiently colonize the human gastrointestinal tract. However, this idea conflicts with reports demonstrating that virulence as tested in a murine model is more frequent in clinical isolates and is associated with particular patterns generated with a restriction fragment length

Molecular Detection of Foodborne Pathogens

polymorphism (RFLP) typing method.56 A study on 118 S. cerevisiae strains from De Llanos et al.53 confirmed that a phenotypic trait fundamental for virulence is the maximum growth temperature: it was observed that the ability to grow at temperature as high as 42°C belongs to different clinical isolates and to very few of the industrial strains: in general these were unable to grow at 42°C, with the exception of “S. boulardii”, the wine strain Uvaferm 71B and the two baker’s yeasts. A gene implicated in the capacity to grow at 42°C is SSD1, which encodes a protein that affects various cellular processes (including maintenance of cell integrity, control of cell cycle, and growth at high temperature), and seems to play an important role in the virulence of Saccharomyces organisms. Deletion of this gene led to a significant increase in virulence for both clinical and plant isolates, as revealed in a mouse model of invasive infection. Alteration in composition and cell wall architecture is supposed to lead to over stimulation of the pro-inflammatory response.64 McCusker et al.63 observed for colonies grown on casein that many of clinical and nonclinical strains formed pseudohyphae (Psh ± phenotype). Pseudohyphal formation was seen more frequently, and tended to be more extensive (longer pseudohyphal chains and more pseudohyphal chains emanating from a colony), for clinical isolates but was not limited to these strains (13 of 15 clinical isolates formed pseudohyphae at 37°C, vs. six of ten nonclinical isolates). Also de Llanos et al.53 evaluated the ability to form pseudohyphae observing that the main potentially virulent isolates expressed this character at much higher levels than the industrial strains, that exhibited low or null levels. The exceptions among the industrial strains were “S. boulardii,” the two baker’s yeasts and one wine strain, Fermivin Crio. In accordance with genetic traits that are implicated in virulence of fungal pathogens,62 de Llanos et al.53 also evaluated the production of extracellular lytic enzymes. Regarding phospholipase activity again a difference was apparent between the groups of clinical and industrial strains: in general the potentially virulent strains produced higher amounts of this enzyme than the industrial strains. An additional virulence factor identified in S. cerevisiae is the heat shock protein hsp90, that when over-expressed determines an increase of the number of colonies in liver, kidney, and in spleen and mortality rate in mice.65 From these evidences it is clear that S. cerevisiae should be considered as an opportunistic pathogen of low virulence rather than a nonpathogenic yeast. Moreover the increasing importance of S. cerevisiae as infective agent creates a need for the development of strain specific molecular markers able to discriminate the potentially pathogenic strains also to avoid unjustified alarm around industrial and foodborne strains. Thus, the possibility to distinguish between potentially pathogenic and harmless strains on the basis of genetic traits is of notable importance. To date molecular methods have been applied mostly in the fingerprinting of pathogenic, clinical or food isolates but no molecular tests have been directed to the detection of genetic traits directly involved in pathogenicity. The typing methods currently available are

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Saccharomyces

merely experimental and all must still be validated for use in infection diagnosis. Moreover, while they are highly discriminant for strain distinction they do not give indications on the potential pathogenicity of the isolates. For example, based on microsatellite polymorphism, it was not always possible to distinguish a clade comprising clinical isolates. Also, there was no separation between strains isolated from blood and cases of vaginitis and those isolated from feces as probably commensal organisms. However, a positive aspect of the application of genotyping methods is the possibility of strain discrimination. As an example, in a study based on microsatellite polymorphisms clinical strains collected from patients not treated with “S. boulardii” exhibited a distinct pattern, with the exception of isolates collected from the same patients and a cluster of four isolates collected from French patients that shared their pattern with that of a commercial baker’s yeast. In this case it was postulated that the colonization of the patients resulted from the use of this strain in the diet.34 Many of the genetic typing methods must be adapted to clinical samples from procedures primarily conceived for food analyses aimed at the definition of biodiversity of natural wine associated strains or of population dynamics during the fermentation processes. However, it is still important to apply methods optimized for different types of food samples in epidemiological investigations, since the implication of bakery yeast strains has been demonstrated in some cases of vaginitis50,51 and in the study of de Llanos et al.58 commercial baker’s strains were indistinguishable from clinical isolates, further showing that the linkage of strains from this food source to human infections is of particular importance. Identification at the strain level is needed to better elucidate the epidemiology of S. cerevisiae infections and to actuate measures for avoiding the transmission of hazardous strains from food. The development, improvement, and validation of molecular markers for identification of S. cerevisiae at the strain level is also important for a better understanding of infections caused by this yeast. For example, in case of relapse, the typing method could be used to trace isolates, to test whether the infection is due to the persistence of the microorganism or reinfection with a new strain. Also, since the risk of transmission of S. cerevisiae from patient to patient has been suggested, it is important to identify isolates in order to demonstrate the origin of the infection and further apply appropriate prevention measures. Molecular techniques for the detection, identification, and typing of potentially pathogenic strains of S. cerevisiae must consist of multistep procedures capable of responding to requisites like the possibility of rapid culture independent detection, secure identification and discrimination from taxonomically related species, primarily those grouped in the S. sensu stricto complex, and strain specific characterization for avoiding use of possibly virulent strains in the food and beverage industry. Methods for executing sequentially these steps have been made available by different experimental studies and can be combined in order to form a work plan able to provide complete analytical evaluation of risks associated to the occurrence of this yeast in clinical and nonclinical

environments and, ultimately, on the epidemiology of infections caused by strains possessing traits of virulence and invasiveness. The following points are of special interest: (i) prompt diagnosis by correct identification of the infecting yeast by a specific culture independent method; (ii) in case of positive results, application of a highly discriminating strain typing method enabling the definition of relationships between isolates from infection cases and strains from sources proximate to the patients; (iii) characterization of the pathogenic potential of foodborne strains by available genetic tests directed to genes involved in virulence and invasiveness. Indeed, feral, and clinical strains of S. cerevisiae showed similar virulence in a mouse model of infection and it is not yet possible to identify particular features of S. cerevisiae that distinguish between environmental and clinical strains on the basis of virulence-related phenotype.63

44.1.5 Molecular Identification and Characterization of S. cerevisiae Strains Molecular techniques commonly used for the suitable identification of isolates of S. cerevisiae as well as other yeast species comprise: the nucleotide sequences of the domains D1 and D2 located at the 5′ end of gene 26S66 and PCR amplification of the ITS1-5.8S-ITS2 rDNA internal transcribed spacer region, followed by digestion with frequent cut restriction endonucleases CfoI, HaeIII, and HinfI.67,68 These techniques are more reproducible and faster that the conventional methods based on physiological and morphological characteristics. Culture-independent molecular techniques based on specific diagnostic sequences have also been developed to detect-quantify S. cerevisiae in wine69 and in feces.70 The latter study applied the direct detection/quantification of S. cerevisiae on nucleic acids extracted from clinical samples. The application of this method could greatly aid the definition of how stably S. cerevisiae strains can inhabit the human gastro-intestinal tract to elucidate the unclear point of whether this microorganism is a commensal or an occasional contaminant. Recently, a specific probe for S. cerevisiae was included in an APEX (arrayed primer extension) microarray support projected for one step detection of common pathogenic fungi based on their ITS1 and ITS2 sequences. Authors stated that identification by this method is unambiguous, sensitive, and reproducible.71 The molecular methods that allow S. cerevisiae strain distinction include karyotyping by pulsed field gel electrophoresis (PFGE), once considered the reference method72 and RFLP analysis on mitochondrial or nuclear DNA.56,73,74 Alternatively, much more rapid PCR based protocols have been developed and applied in studies regarding the distinction of food or clinical isolates, such as the tests directed to the intron conserved sequence motifs,75 to the transposon δ elements,74,76,77 and to the microsatellite33,35,78 and minisatellite repeats79 of different gene loci. For intron targeted

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Molecular Detection of Foodborne Pathogens

PCR, applied for the distinction of commercial strains, primers were designed on the conserved spliceosome recognition sequences with a 3′ end complementary to the 5′ splice site and to the lariat branch consensus. The primers were extended at their 5′ end to produce 16-mer oligonucleotides by using random sequence. These were used singularly or in pair with best results of strain discrimination when primers corresponding to both consensus sequences were used in combination.75 The method exploited the high level of polymorphism present in the yeast intron sequences, that are not under the action of evolutionary constraints. Recently, PCR amplification with primers targeted on the retrotransposon δ sequences (interdelta sequence typing) has been improved,77 and can be considered as discriminanting as karyotyping or mtDNA restriction analysis.80 Amplifications with the improved primers designed by Le Jeune et al.81 generated more polymorphic patterns than amplification with the basic primers of Ness et al.76 that were not influenced by the amount of template DNA. According to the guidelines proposed by the European Study Group on Epidemiological Markers,48 it was found that the application of microsatellite polymorphism to S. cerevisiae is a powerful method of strain distinction. Microsatellites are short sequence repeats (SSR), usually less than ten nucleotides and highly polymorphic in length found in many open reading frames (ORF) and in noncoding regulatory regions. All the S. cerevisiae strains, including “S. boulardii,” gave 100% positive PCR results with all microsatellites tested.33 In contrast, isolates belonging to the closely related species S. pastorianus, S. paradoxus, and S. bayanus failed to be reproducibly amplified limitedly to the loci considered, therefore the method was proven to be specific for S. cerevisiae.33 Among the molecular typing techniques the profiles of microsatellites amplification from different loci was found to be highly reproducible, able to discriminate strains from different sources, as well as “S. boulardii”

M

1

2

3

4

isolates from treated patients and from pharmaceutical preparations, and exploitable for phylogenetic clustering. In the work of Malgoire et al.35 reproducibility of the method and stability of the genomic target regions were confirmed. The level of discrimination between strains obtained was never previously reached with other methods applied to S. cerevisiae typing, confirming the high degree of polymorphism of microsatellite regions among strains of S. cerevisiae. In the experiments of Hennequin et al.33 and Posteraro et al.82 an allele of locus YLR177w, (CAG)9, was found specific for “S. boulardii” by this technique. Another important advantage of the method is its portability. Since all results can be expressed as a number of repeats, computer translation is easy, offering the opportunity to compare results from different groups via the Internet. This is an important issue for further comparisons of larger sets of strains isolated in very different settings, e.g., human medicine, environmental studies, and wine production. Also it should be underlined that the technique can be used with a nonradioactive labelling and separating the amplification products on agarose rather than polyacrilamide gel, according to recent modifications of this method.83 Examples of amplification patterns obtained as described by Vaudano and Garcia-Moruno83 are reported in Figure 44.1. The presence of repeat sequences inside the ORFs of cell wall protein genes is very common and well-documented in S. cerevisiae and other yeasts. These minisatellite repeats are ricombinogenic and can be a source of genetic variability among strains. The kind of minisatellite selected for wine strain typing by Marinangeli et al.79 within sequences available for the reference strain S. cerevisiae S288c at http://www. yeastgenome.org satisfied the following criteria in order to allow amplification from different strains and detection on agarose gel; i.e., at least 80% identity between repeats and at least 20 bp repeat length, respectively. Moreover the test was restricted to structural nonenzymatic proteins because

5

6

7

8

M

Figure 44.1  Examples of band patterns generated by the microsatellite amplification method of Vaudano and Garcia-Moruno. Lines M, 1 kb plus DNA ladder (Invitrogen); lines 1–8, S. cerevisiae isolates from different wineries.

Saccharomyces

they were supposed to be more variable since their sequence variation would not cause a lethal loss of function. Four genes were selected: AGA1 coding for the anchoring subunit of α-agglutinin, DAN4, member of the PAU proteins of unknown function and HSP150, a cell-wall bound heat shock protein and the major cell wall glycoprotein SED1. In Table 44.1 all the mostly adopted S. cerevisiae typing methods are listed and briefly described.

44.2  Methods 44.2.1 Reagents and Equipment Considerable research efforts have been directed toward the development of sensitive and specific tools for the identification of S. cerevisiae isolates. In this section we provide our recommendations for test methods that are suitable for detection and typing of S. cerevisiae isolates from different matrices. Table 44.2 lists the reagents and equipment needed for the analysis relative to the sample treatment and subsequent identification of S. cerevisiae that will be described in detail in the following paragraphs.

44.2.2 Sample Treatment Most studies on the genetic typing of clinical S. cerevisiae strains were carried out on isolates obtained by hospitals or medical centres after isolation by culturing methods applied in routine diagnosis.33,35,52,58 The strains had been received by the research laboratories already physiologically identified as S. cerevisiae by standard automated systems such as API 20C, API ID 32C and Vitek 32 (BioMérieux, Marcy l’Etoile, France). The clinical laboratory routine cultures from which isolates were collected were from specimens like blood, stools, urine, sputum or oral swabs, vaginal discharges, or swabs and bronchoalveolar lavages, given the wide variety of infectious states that can be caused by virulent S. cerevisiae strains. The sample collection and analysis procedures varies for each material drawn: in the case of blood cultures the sample is transferred directly from the patient suspected of fungal infection to bottles containing the appropriate growth medium for a separate fungal culture and soon incubated. Similarly swabs are immediately transferred into tubes of fungi specific cultural broth and incubated. Feces are collected as random sample in sterile containers and later submitted to standard homogenization, dilution, and plating procedures. The isolation of yeasts is currently performed on Sabouraud dextrose agar (10 g/l mycological peptone, 40 g/l dextrose, pH 5.6) plus 4 g/l chloramphenicol. The decision on whether the isolated yeast is a contaminant or a pathogen is done on the basis of patient history, physical examination, body temperature, clinical course, and laboratory data (i.e., culture results, white blood cell count). Hence, the characterization of each yeast isolate by genetic methods can be executed after correct assignment of the strains under study to the categories of pathogenic or of contaminant saprophyte.

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(i) Extraction of DNA from pure cultures of S. cerevisiae: For preparing pure cultures of the isolated clinical S. cerevisiae strains single colonies, picked from the isolation plates containing 30–300 colonies and chosen according to morphological features like shape, color, size, are streaked on the surface of yeast specific agar plates. After growth, a well isolated colony is transferred and allowed to develop in an agar slant or in broth for the preparation of refrigerated or frozen cultures for short and long term storage, respectively. The refrigerated slants can be utilized within two weeks. The cultures for long term storage are habitually frozen after addition of 20% v/v glycerol in sterile 2 ml cryovials and kept at –20°C or –80°C for one or two years or for several years, respectively. The preparation of lyophilized material is required for storing the strains for up to 20 years. To this aim the yeast culture is freeze dried on a cryoprotectant, e.g., skim milk, and sealed in a glass ampoule. At the moment of use the yeast is inoculated from a frozen culture by scraping the surface and transferring the inoculum into a slant or broth culture. The rest of the culture is kept frozen and stored again. DNA suitable for typing by a PCR based method from a pure culture of S. cerevisiae can be derived from the protocol of Hennequin et al.33 applied prior to microsatellites amplification. The required reagents, consumables, and equipment are listed in Table 44.2. The procedure includes the following steps: (1) Grow the S. cerevisiae isolate overnight at 30°C in 3 ml of YPGlu medium. (2) Harvest yeast cells from 3 ml of culture by centrifugation at 10,000 × g for 5 min. (3) Remove the supernatant and wash the cell pellet with 1 ml of TE buffer; centrifuge the cells at 10,000 × g for 5 min. (4) Resuspend the cell pellet in 200 µl of lysis buffer; incubate for 1 h at 37°C. (5) Add 200 µl of lysing solution; incubate at 65°C for 30 min; keeping on ice for 5 min. (6) Add 100 µl of potassium acetate 5 M; keeping at 4°C for 45 min; centrifuge for 5 min at 10,000 × g. (7) Transfer the supernatant in a clean tube; precipitate DNA by addition of 30 µl of ammonium acetate solution plus 3 volumes of 100% ethanol; centrifuge for 10 min at 10,000 × g. (8) Remove the supernatant and add 1 ml of 70% ethanol; centrifuge for 2 min at 10,000 × g. (9) Carefully remove the ethanol by gentle aspiration; dry in the open tubes under a fume cabinet until all residual ethanol evaporates. (10) Resuspend in 50 µl of TE buffer; store at 4°C for up to 6 months. (ii) Extraction of total DNA from human fecal samples: The application of direct detection of S. cerevisiae in clinical samples by specific PCR would expedite the diagnosis of infections due to this yeast and would decrease the possibility of misidentification by physiological assays. The

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Molecular Detection of Foodborne Pathogens

Table 44.1 Molecular Methods Developed for the Discrimination of S. cerevisiae Isolates at the Species or Strain Level Method Karyotyping

EcoRI RFLP

mtDNA restriction

Intron splice site targeted PCR

ITS1-5.8S-ITS2 region amplification and restriction

Amplification of δ sequences

Amplification of microsatellite sequences

Amplification of minisatellites in cell wall protein genes

Real-time PCR targeted on rRNA genes and ITS

Steps of the Procedure Purification of total DNA by extraction from cells immobilized in agarose plugs Electrophoretic separation in a minimum of 23 h by PFGE Profile analysis. Total DNA extraction from pure cultures Digestion for 90 min with EcoRI endonuclease 21 h run on 1% agarose gel Profile analysis Total DNA extraction from pure cultures Digestion overnight with HinfI or RsaI endonucleases Run on 1 % agarose gel Profile analysis Total DNA extraction from pure cultures by freeze boil treatment of cells Amplification with one of two primer sets designed on intron conserved sequence motifs (each primer can be used singularly) Separation on 2% agarose gel Profile comparison DNA extraction from pure cultures PCR with primers flanking the ITS1-5.8SITS2 region Restriction with endonucleases CfoI, HaeIII, and HinfI Separation on 3% agarose gel DNA extraction from pure cultures PCR with primers targeted to transposon δ sites separation on 1.5% agarose gel profile comparison DNA extraction or cell disruption by freeze-thaw treatment from pure cultures PCR with primers flanking microsatellite sequences of different genetic loci Separation on polyacrylamide gels containing formamide or on agarose gel Profile comparison DNA extraction or cell disruption by heat treatment from pure cultures PCR amplification with a primer set targeted on the sequences flanking the minisatellite regions of cell-wall protein genes Separation on agarose gel Profile comparison DNA extraction by mechanical cell disruption Amplification in a normal PCR reaction followed by conventional electrophoresis or in a real-time PCR apparatus for quantification using Sybr Green as a fluorescent dye

Discrimination Level

Disadvantages

Single strain

Time consuming Delicate and expensive equipment

Strain clusters

High amount of unfragmented DNA needed Time consuming No distinction of the restriction profile group B from S. paradoxus

Single strain

Relatively high amount of unfragmented DNA needed for digestion (~0.5 µg) Time consuming

Single strain

To date experimented only on few commercial strains

Species

None

Single strain

Unstable patterns with some primer sets Not all strains produce amplification products

Single strain

Need of separation on polyacrylamide gels in a sequencing apparatus in the original method or labeling with different fluorochromes due to the narrow range of dimension of the amplification products obtained for some genetic loci

Single strain

Need to combine the profiles obtained for amplicons from different loci with four different primer pairs

Culture-independent detection and identification of S. cerevisiae

To date experimented only on feces and wine

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Table 44.2 List of Materials Needed for Different Molecular Analyses on S. cerevisiae Equipment

Reagents

A. DNA Extraction from Pure Cultures of S. cerevisiae YPGlu medium (10 g/l yeast extract, 10 g/l Disposable gloves (latex, vinile or nitrile) peptone, 20 g/l glucose) Eppendorf-like sterilized 2 ml micro tubes TE buffer (10 mM Tris/HCl, 1 mM EDTA, pH 8.0) Sterilized pipette tips fitting to Gilson Lysis buffer (50 mM Tris-HCl pH 8, 25 mM P20/P200, P1000 EDTA, 1% vol/vol β-mercaptoethanol, 100,000 U zymolyase) Lysing solution (200 mM diethanolamine, pH 9.0, 80 mM EDTA, pH 9.0, and sodium dodecyl sulfate 1 % wt/vol) Potassium acetate 5 M Ammonium acetate 25 M Absolute ethanol 70 % ethanol

Culture incubator Benchtop centrifuge Variable volume micropipettes corresponding to Gilson P200 and P1000

80°C freezer Benchtop refrigerated centrifuge Bunsen burner Sterile air flow cabinet Technical balance Liquid nitrogen dispenser tank Variable volume micropipettes corresponding to Gilson P10, P200, and P1000

Labware and Consumables

B. Extraction of Total DNA from Human Fecal Samples Extraction buffer (1% wt/vol, sarkosyl 100 mM Disposable gloves (latex, vinile or nitrile) Tris-HCl, 100 M NaCl, 10 mM, EDTA, 2% Sterile sampling bags wt/vol, polyvinylpolypyrrolidone, 0.1 M Sterilized ceramic mortars and pestles Na2SO3, pH 8.0) Metal spoon flame sterilizable spatula Phenol–chloroform–isoamyl alcohol (25:24:1) 2 ml Eppendorf-like sterilized safe lock micro vol/vol/vol mixture tubes 3 M NaAc pH 4.8 Sterilized pipette tips fitting to Gilson Isopropanol P20/P200, P1000 70% ethanol TE buffer (10 mM Tris/HCl, 1 mM EDTA, pH 8.0) DNase free RNase A

C. Microbial Cells Separation Prior to Total DNA Extraction from Bakery Dough Stomacher homogenizer PBI 400 Sterile saline-tryptone diluent (0.9% w/v NaCl, Disposable gloves (latex, vinile or nitrile) Pipetting device (Pipetus or equivalent) 1% w/v casein tryptone) Stomacher bags Bunsen burner Sterile 25 ml plastic pipettes High capacity refrigerated centrifuge equipped Sterilized polypropylene 50 ml centrifuge with 50 ml tubes fitting rotor tubes Nalgene or, in alternative, 50 ml Refrigerated benchtop centrifuge Falcon-like tubes –20°C freezer

High capacity refrigerated centrifuge equipped with 10 ml tubes fit rotor adapter Pipetting device (Pipetus or equivalent) Variable volume micropipettes corresponding to Gilson P200 and P1000 Mini Bead Beater (Biospec, OK)

Quantitative PCR apparatus, e.g., OPTICON™2 DNA Engine (MJ Research) or GeneAmp 5700 sequence detection system (PE Applied Biosystems) PCR dedicated flow cabinet (optional) Variable volume micropipettes corresponding to Gilson P2, P20, P200, and P1000

D. Extraction of total DNA from wine Sterile water PrepMan kit (PE Applied Biosystems) Absolute ethanol 70% ethanol

E. Detection of S. cerevisiae by Real-Time PCR SYBR green PCR ready master mix, e.g., DyNAmo™ HS SYBR® qPCR kit (FINNZYMES) or SYBR green PCR master mix (PE Applied Biosystems) Specific primers pairs Sterile water

Disposable gloves (latex, vinile or nitrile) Sterile 10 ml plastic pipettes Sterilized polypropylene 50 ml centrifuge tubes Nalgene or, in alternative, 50 ml Falcon-like tubes Sterilized pipette tips for Gilson P2/P10, P20/P200, P1000 Sterilized 2 ml plastic tubes (Eppendorf) Sterilized zirconia/silica 0.5 mm beads

Disposable gloves (latex, vinile or nitrile) Sterile pipette tips for Gilson P2/P10, P20/P200, P1000 with filter 0.2 ml thin wall sterile tubes or tube strips with optical caps for fluorescence detection

(Continued)

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Table 44.2  (Continued) Equipment

Reagents

Labware and Consumables

F. Identification of S. cerevisiae by Restriction of the Amplified 5.8S-ITS Region of the Ribosomal rRNA Operon Disposable gloves (latex, vinile or nitrile) Thermal cycler Taq DNA polymerase Sterile pipette tips for Gilson P2/P10, P20/P200, PCR dedicated flow cabinet (optional) PCR reaction buffer (100 mM Tris/HCl, pH 8.3 P1000 with filter Variable volume micropipettes corresponding at 25°C; 500 mM KCl; 0.01% gelatin), usually 0.5 or 0.2 ml thin wall sterile tubes with caps to Gilson P2, P20, P200, and P1000 supplied with the enzyme Horizontal electrophoresis system and power 25 mM MgCl2 solution Deoxinucleotide (dNTPs: dATP, dGTP, dCTP, supply dTTP) mix stock solution ITS1/ITS4 primers stock solutions Sterile water Sterilized mineral oil Endonucleases CfoI, HaeIII, and HinfI 1 × Tris acetate EDTA (TAE) buffer (4.8 g/l Tris base, 1.14 ml/l acetic acid, 0.29 g/l EDTA, pH 8.1) Agarose for routine use Ethidium bromide, 50 mg/ml aqueous solution

Thermal cycler PCR dedicated flow cabinet (optional) Variable volume micropipettes corresponding to Gilson P2, P20, P200, and P1000 Horizontal electrophoresis system and power supply

G. Typing of S. cerevisiae by Microsatellite Amplification Disposable gloves (latex, vinile or nitrile) Taq DNA polymerase Sterile pipette tips for Gilson P2/P10, P20/P200, PCR reaction buffer (100 mM Tris/HCl, pH 8.3 P1000 with filter at 25°C; 500 mM KCl; 0.01% gelatin), usually 0.5 or 0.2 ml thin wall sterile tubes with caps supplied with the enzyme 25 mM MgCl2 solution Deoxinucleotide (dNTPs: dATP, dGTP, dCTP, dTTP) mix stock solution Multiplex primer set flanking microsatellites at different loci Sterile water Sterilized mineral oil Agarose for routine use 0.5 × Tris borate EDTA (TBE) buffer (5.4 g/l Tris base, 2.75 g/l boric acid, 2.92 g/l EDTA, pH 8.1) Ethidium bromide, 50 mg/ml aqueous solution

efficiency of DNA extraction is a critical element for sensitivity and repeatability of the molecular typing methods; yield and quality of the genetic material must be at the maximum in order to achieve sensitivity and elimination of false negatives. This aspect has even more prominence in quantitative detection protocols. The crucial step in the yield of genetic material from yeasts, regardless if from a sample or pure culture, is cell lysis. Instead, the prosecution of purification can be easily and reproducibly accomplished by a number of available commercial kits. A study focused on the comparison of fungal cell wall disruption methods evaluated by optical microscopy the efficiency of breaking hyphae, conidia or cells with chemical, physical or mechanical treatments, on 16 fungal pathogens, including a strain of Candida albicans as representative of the yeast species.84 To efficiently extract a high amount of good quality DNA from all the species considered it was necessary to combine the enzymatic treatment with mechanical disruption by mortar and pestle grinding of material frozen in liquid nitrogen. This method was applied later by another research

group and allowed high sensitivity in the PCR based detection of S. cerevisiae and S. sensu stricto DNA from feces.70 In the reported experiments, fecal samples were drawn from 11 healthy adults and were stored at –80°C in sterile plastic bags before DNA extraction. The list of reagents and equipment recommended are reported in Table 44.2. The phases of the method are as follows: (1) Thaw the fecal sample; transfer about 1 g of it in a laboratory mortar previously well cleaned and sterilized by keeping overnight at 200°C in a stove. (2) Add 500 µl of extraction buffer; immediately freeze by liquid nitrogen insufflations. (3) Grind for 2 min; add 500 µl of phenol-­chlorophormisoamyl alcohol mixture and rotary mixing at room temperature for 10 min. (4) Transfer the sample to a 2-ml centrifuge tube; centrifuge at 10,000 × g for 10 min at 4°C. (5) Carefully transfer the supernatant in a new tube.

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Saccharomyces

(6) Add 500 µl isopropanol and 150 µl of 3 M sodium acetate solution pH 4.8; mix for 5 min and keep for additional 5 min at room temperature; centrifuge at 10,000 × g for 10 min at room temperature. (7) Carefully discard the supernatant and add 500 µl 70% ethanol to the pellet; centrifuge for 2 min at 10,000 × g. (8) Remove as much as possible ethanol by gentle aspiration; dry in the open tubes under a fume cabinet until all residual ethanol evaporates. (9) Resuspend in 100 µl of TE buffer containing DNase free RNase A; store at 4°C allowing RNA digestion overnight. (iii) Microbial cells separation prior to total DNA extraction from bakery dough: The types of food that can contain high numbers of S. cerevisiae, and therefore are worth of being preferentially taken into account for epidemiological studies, are wines and bakery doughs. The main scope is of defining the occurrence of potentially pathogenic S. cerevisiae strains in these food sources. Thus, the clarification of whether they are implicated in the epidemiology of S. cerevisiae infections must rely on strain isolation and characterization. One example of medium frequently used for yeast isolation from these food materials is YPD (yeast extract 10 g/l, peptone 10 g/l, glucose 20 g/l) supplemented with 0.1 g/l chloramphenicol or 0.015 g/l biphenyl to inhibit bacteria and moulds, respectively. Sample treatments applied for the separation of yeast cells from wine and bakery doughs or starters, were assessed to follow natural yeast dynamics during food production, but their application can be extended to the evaluation of S. cerevisiae related risk. A method used by Meroth et al.85,86 to separate microbial cells from a bread dough in order to study the population dynamics during fermentation by PCR-DGGE comprises earlier sample treatment steps that can be carried out also for the isolation of yeast strains from the baker’s dough. Materials used are listed in Table 44.2. The operating processes are as follows: (1) Draw about 10 g of sample by a sterile spoon spatula and transfer it in a stomacher bag (at this stage the sample can be stored frozen at –20°C if it must not be analyzed soon). (2) Add 90 ml of saline-tryptone diluent each 10 g of sample; homogenize for 5 min in stomacher. (3) Centrifuge a 50 ml aliquot at 4°C for 5 min at 200 × g or for 5 min or at 1,500 × g, depending on the tendency to sedimentation of the suspended solid particles. (4) Directly cultivate by dilution and plating. Alternatively, concentrate the cells by centrifugation for 15 min at 5,000 × g in order to isolate also the less abundant microbiota components. The cell pellet can be stored at –20°C if it will be used for DNA extraction. (iv) Extraction of total DNA from wine: The culture independent detection of S. cerevisiae in this kind of food

sample, either by PCR-DGGE targeted on 28S rRNA gene or by species-specific PCR is of interest for the individuation of the samples or the sample stage at which this yeast predominates, given that the variability of S. cerevisiae number and occurrence in bread fermentation batches is high.86 DNA isolation from the cell pellets prepared as described can follow the same procedure reported above for the pure cultures. The isolation of yeasts from wine or other fermented beverages is accomplished by direct dilution and plating during fermentation or concentrating by centrifugation in sterile tubes before plating on suitable cultural media at the end of fermentation or after stabilization of the final product. Indeed yeast counts during wine storage are expected to be null or very low. The materials needed according to Martorel et al.69 are reported in Table 44.2. The phases of sample treatment and analysis are as follows: (1) Draw 10 ml sample from bottles or bags opened in a sterile environment by a sterile pipette or transfer 10 ml of the wine in a sterile container directly from the barrel or winery vat; keep refrigerated until analysis or until freeze at –20°C. (2) Plate the diluted sample from a product in course of fermentation, or concentrate by centrifugation at 10,000 × g for 5 min of the stabilized wine and use the concentrated pellet for yeast isolation. In case it is preferred to explore the presence and number of S. cerevisiae before undertaking isolation, a method for the direct detection of this yeast in wine down to 3.8–5 CFU/ml by quantitative real-time PCR69 can be adopted. The protocol comprises the following steps: (1) Centrifuge 10 ml authentic wine sample 10,000 × g for 5 min. (2) Wash the pellet twice with sterile water. (3) Physically disrupt the cells resuspended in PrepMan buffer in a mini-bead beater after adding 0.3 g of 0.5-mm zirconia/silica beads in three repetitions of 30 sec at high speed and cooling samples on ice for 1 min between the breakdown steps. (4) Extract DNA according to the PrepMan kit manufacturer’s protocol. (5) Precipitate with two volumes of absolute ethanol to minimize inhibitions; centrifuge at 10,000 × g for 10 min. (6) Wash with one volume of 70 % ethanol; centrifuge and carefully remove the ethanol by aspiration. (7) Dry the pellet by exposure to air under a fume cabinet; resuspend DNA in 50 µl of sterile water.

44.2.3  Detection Procedures 44.2.3.1 Detection of S. cerevisiae by Real-time PCR The specific identification and quantification of S. ­cerevisiae by PCR method is possible by two protocols for the

632

detection-quantification of this yeast directly in feces70 and in wine,69 respectively, by quantitative real-time PCR (qPCR) using the SYBR green fluorochrome. (i) Detection of S. cerevisiae in feces: The oligonucleotides SCDF (5′-AGGAGTGCGGTTCTTTG-3′ and SCDR (5′-TACTTACCGAGGCAAGCTACA-3′) are used. These primers target the D1/D2 region of 26S rRNA gene and amplify a 310-bp DNA fragment from S. cerevisiae, but not from other S. sensu stricto species. The materials needed are listed in Table 44.2. Amplification is done using DyNAmo™ HS SYBR® qPCR kit (FINNZYMES) master mix, 10 pmol each primer and 1 µl template prepared according to the method of DNA extraction from feces described above in 20 µl of PCR mixture. The cycling program comprises initial denaturation at 95°C for 15 min, 45 cycles of denaturation at 94°C for 20 sec, annealing at 58°C for 30 sec and extension at 72°C for 45 sec and final extension at 72°C for 5 min. Amplification is performed in an OPTICON™2 DNA Engine, MJ Research. (ii) Detection of S. cerevisiae in wines: The oligonucleotide pair used in qPCR from wine69 are: SC1d (5′-ACA TATGAAGTATGTTTCTATATAACGGGTG-3′) and SC1r (5′-TGGTGCTGGTGCGGATCTA-3′), which are designed on a RAPD PCR band sequence obtained only from S. cerevisiae strains, and enable discrimination of S. cerevisiae from its sibling species S. bayanus, S. pastorianus and S. paradoxus. These primers allow the amplification of a 301-bp product from S. cerevisiae also in presence of contaminant yeasts and bacteria. The PCR mixture contains 12.5 µl of SYBR green PCR master mix (PE Applied Biosystems), 2.5 µl of DNA at 10 ng/ml, and 60 mM each primer in a 25 µl reaction. To minimize inhibition by substances present in the culture medium or in the wine, bovine serum albumin (BSA) is added to the PCR mixture in a final concentration of 0.05 µg/µl. The PCR program is: 95°C for 10 min and 40 cycles of 95°C for 15 sec, 60°C for 1 min, and 72°C for 1.5 min. Amplification is carried out in a GeneAmp 5700 sequence detection system (PE Applied Biosystems). 44.2.3.2 Identification of S. cerevisiae by Restriction of the Amplified 5.8S-ITS Region of the Ribosomal rRNA Operon When DNA from pure cultures is available, a rapid method able to identify S. cerevisiae is PCR amplification of the ITS15.8S-ITS2 rDNA internal transcribed spacer region, followed by digestion with frequent cut restriction endonucleases67,68 and by separation on a 3% agarose gel. For this purpose oligonucleotides ITS1 (5′-TCCGTAGGTGAACCTGCGG-3′), and ITS4 (5′-TCCTCCGCTTATTGATATGC-3′), delimiting a region of the rRNA coding repeat unit comprising the 5.8 S rRNA gene and the two internal transcribed spacers ITS1 and ITS2, are used. The profile corresponding to S. cerevisiae is distinct from those corresponding to the other S. sensu stricto species and shows bands of the following molecular weights obtained by restriction of a 880-bp amplicon: 385 bp and 365 bp with CfoI, 320 bp, 230 bp, 180 bp and 150 bp with

Molecular Detection of Foodborne Pathogens

HaeIII and 365 bp and 155 bp with HinfI. This amplification/digestion procedure is applicable also directly to isolated yeast colonies for rapid identification and comprises the following steps using the materials listed in Table 44.2. (1) Prepare PCR mixture containing 0.5 µM each primer, 10 µM deoxynucleotides, 1.5 mM MgCl2 and 1 × PCR buffer, sterile double-distilled water up to the desired volume and 1 µl of DNA sample in a 10 µl reaction. In the original method of direct amplification from a single colony the cell material is added to a reaction mixture of 100 µl final volume and 1 U of Taq DNA polymerase is added after heating of the mixture containing the cell material for 15 min at 95°C. (2) The cycling program consists of initial denaturation at 95°C for 5 min; 35 cycles of denaturing at 94°C for 1 min, annealing at 55±5°C for 2 min and extension at 72°C for 2 min; and a final extension at 72°C for 10 min. (3) Digest the PCR products (10 µl or approximately 0.5–1.0 µg) without further purification with 1 U/ µl of the restriction endonucleases CfoI, HaeIII, and HinfI at 37°C for 2–3 h (or overnight); (4) Separate the PCR products and their restriction fragments on 1.4% and 3% agarose gels, respectively, in 1 × TAE buffer. (5) Stain the gels in 0.5 µg/ml ethidium bromide aqueous solution. (6) Estimate the sizes of PCR amplicons by comparison against a DNA length standard (e.g., 100 bp ladder, Invitrogen). 44.2.3.3 Typing of S. cerevisiae by Microsatellite Amplification For S. cerevisiae strain distinction, the microsatellite typing method is valuable due to its rapidity, high output and high strain discrimination potential. The procedure described below is based on a previous report83 with materials listed in Table 44.2. Primers flanking microsatellites at loci SC8132X, YOR 267C, and SCPTSY7 are used, their sequences are: SC81 32XFW (5′-CTGCTCAACTTGTGATGGGTTTTGG-3′) and SC8132XRV (5′-CCTCGTTACTATCGTCTTCATCTTGC-3′), YOR267CFW (5′-GGTGACTCTAACGGCAGAGTGG-3′) and YOR267CRV (5′-GGATCTACTTGCAGTATACGGG-3′), SCP TSY7FW (5′-AAAAGCGTAAGCAATGGTGTAGAT-3′) and SCPTSY7RV (5′-AAATGATGCCAATATTGAAAAGGT-3′). The 20 µl PCR mixture is made up of 3.1 mM MgCl2, 400 µM dNTPs, 2 U of Taq DNA polymerase (Sigma Co.), 1 µl of DNA and 10, 15, and 40 pmol of the YOR267C, SC8132X, and SCPTSY7 targeted primer sets, respectively. The PCR amplification is run in a MyCycler machine (BioRad) with the following steps: initial denaturation for 4 min at 94°C, 28 cycles of denaturation at 94°C for 30 sec, annealing at 56°C for 45 sec, elongation at 72°C for 30 sec,

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Saccharomyces

final extension at 72°C for 10 min. The amplification products are separated by running on a 2.5% w/v agarose gel in 0.5 × TBE buffer (5.4 g/l Tris base, 2.75 g/l boric acid, 2.92 g/l EDTA pH 8.1) at 100 V for 80 min and stained with 0.5 µg/ml ethidium bromide.

44.3 Conclusions and Future Perspectives Future investigations must improve the knowledge of how strains of S. cerevisiae cause disease and, most importantly, how they can be controlled. Direct detection of S. cerevisiae from clinical samples could be extremely useful in prompt diagnosis, given some cases of retarded treatment with fungicidal drugs.87 This can be done in analogy with what has been carried out for Candida species in a study regarding the detection of yeasts directly from culture positive blood samples from which DNA suitable for LightCycler amplification was simply obtained by adding yeast-positive blood culture medium to an equal volume of lysis buffer containing guanidine hydrochloride. Successful amplification was accomplished on the nucleic acids purified by organic solvent extraction and ethanol precipitation.88 Also it will be important to validate the APEX oligonucleotide microarray support developed by Campa et al.71 for routine practice, indeed it could represent a powerful tool for the rapid and simultaneous diagnosis of S. cerevisiae as well as of many other fungal infections. Use of this analysis tool experimented on pure cultures could be extended to direct detection from clinical specimens. Another aspect to be implemented is the introduction of selection criteria for strains to be used in the food and beverage industry. As suggested by de Llanos,52 these must comprise the evaluation of characters strongly associated to pathogenicity like growth capacity at 42°C, pseudohyphal growth and high levels of phospholipase activity. These selection criteria should be taken into account also for evaluating the risk associated to multi-strain fermentations, for example in winemaking processes in which vineyard and winery indigenous strains are used to improve the sensorial quality and distinctness of the product. Among genetic traits possibly implicated in virulence only some are currently known and their role has still to be defined in natural S. cerevisiae strains. These are the SSD1 gene, whose inactivation causes an increase of virulence due to acquired capacity to grow at 42°C,64 the ammonia transporters MEP genes, that show higher expression during pseudofilamentation,30,89 the gene PLB1 encoding a phospholipase 90 and Hsp90.65 No data are available on the occurrence and expression of these genes in large panels of Saccharomyces isolates and further studies should be carried out to elucidate their contribution to virulence and invasiveness. A similar interest should be devoted to other genetic determinants like those homologous to traits that influence pathogenesis in better known yeast infections like candidiases. Future investigations should be directed at monitoring both the presence and copy number of these genes in food borne

strains compared to clinical and pathogenic isolates, but also to evaluate their level of expression in environmental conditions commonly encountered during food and beverage production, storage, and distribution as well as in the cases in which S. cerevisiae can be found as a microbial contaminant of food products. Moreover, the availability of whole genome sequence data can allow the design of DNA microarray supports applicable to the comparison of the gene sets among highly virulent and non virulent strains; thus other possible virulence target genes to be detected specifically in strains to be evaluated for food and beverage production can be identified. This kind of investigation has been already applied to “S. boulardii” using Affimetrix oligonucleotide chips designed on the whole S. cerevisiae sequenced genome. Comparison with strain S. cerevisiae BY4743, a diploid strain with two copies of each chromosome, permitted to define distinctive genetic features of “S. boulardii,” like the loss of most Ty1/2 elements, trisomy for chromosome IX, and, significantly, an higher copy number for genes of the cAMP pathway involved in pseudohyphal growth. In the study, the enhanced ability of forming pseudohyphae in conditions of low nitrogen availability was confirmed also physiologically. Moreover higher resistance to low pH was observed in “S. boulardii” compared to other S. cerevisiae strains. Other genes found to be overrepresented are involved in protein synthesis and stress response.91 From the results obtained, new DNA microarray supports comprising only genetic determinants found to be distinctive of virulent strains might be designed and applied for the distinction of strains that have an unequivocal pathogenicity potential. Studies on the expression of these genetic traits might be necessary in cases in which the presence of a putative virulence gene is not translated in a virulence phenotype: in this way it is possible to fix the environmental conditions that favor the switch to pathogenicity. The restriction of the analytical procedures to few virulence traits could greatly facilitate the evaluation of strains suitable for food production. In this way a reliable analytical procedure might be made available to assess the safety of strains of S. cerevisiae to be admitted for use in the food industry. Since now the microsatellite polymorphism detection method represents a powerful tool of strain distinction;34,35,83 a standardization of the target loci, PCR, and electrophoresis conditions with creation of a large database comprising clinical isolates could allow the comparison and grouping of new strains with those used as references. Though this will not give information on the pathogenicity of the isolates studied, at least could indicate the possibility to colonize the human body on the basis of genetic similarity with strains from this source.

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Molecular Detection of Foodborne Pathogens 26. Kuthan, M., et al. Domestication of wild Saccharomyces cerevisiae is accompanied by changes in gene expression and colony morphology. Molecular Microbiol., 47, 745, 2003. 27. Palkova, Z. Multicellular microorganisms: laboratory versus nature. EMBO Rep., 5, 470, 2004. 28. Madhani, H.D. and Fink, G.R. The control of filamentous differentiation and virulence in fungi. Trends Cell Biol., 8, 348, 1998. 29. Casalone, E. et al. Characterization of Saccharomyces cerevisiae natural populations for pseudohyphal growth and colony morphology. Res. Microbiol., 156, 191, 2005. 30. Townsend, J.P. and Hartl, D.L. Bayesian analysis of gene expression levels: statistical quantification of relative mRNA level across multiple strains or treatments. Genome Biol., 3, research 0071.1–0071.16, 2002. 31. McCullough, M.J. et al. Species identification and virulence attributes of Saccharomyces boulardii (nom. inval.). J. Clin. Microbiol., 36, 2613, 1998. 32. van der Aa Kühle, A. and Jespersen, L. The taxonomic position of Saccharomyces boulardii as evaluated by sequence analysis of the D1/D2 domain of 26S rDNA, the ITS1-5.8S rDNA-ITS2 region and the mitochondrial cytochrome-c oxidase II gene. Syst. Appl. Microbiol., 26, 564, 2003. 33. Hennequin, C. et al. Microsatellite typing as a new tool for identification of Saccharomyces cerevisiae strains. J. Clin. Microbiol., 39, 551, 2001. 34. Molnar, O. et al. Genotyping identification of Saccharomyces species using random amplified polymorphic DNA analysis. System. Appl. Microbiol., 18, 136, 1995. 35. Malgoire, J.Y. et al. Typing of Saccharomyces cerevisiae clinical strains by using microsatellite sequence polymorphism. J. Clin. Microbiol., 43, 1133, 2005. 36. Billoo, A.G. et al. Role of a probiotic (Saccharomyces boulardii) in management and prevention of diarrhoea. World J. Gastroenterol., 12, 4557, 2006. 37. Sazawal, S. et al. Efficacy of probiotics in prevention of acute diarrhoea: a meta-analysis of masked, randomised, placebocontrolled trials. Lancet Infect. Dis., 6, 374, 2006. 38. McFarland, L.V. Meta-analysis of probiotics for the prevention of traveler’s diarrhea. Travel Med. Infect. Dis., 5, 97, 2007. 39. Whelan, K. Enteral-tube-feeding diarrhoea: manipulating the colonic microbiota with probiotics and probiotics. Proc. Nutr. Soc., 66, 299, 2007. 40. Castagliuolo, I. et al. Saccharomyces boulardii protease inhibits the effects of Clostridium difficile toxins A and B in human colonic mucosa. Infect. Immun., 67, 302, 1999. 41. Buts, J.P. and De Keyser, N. Effects of Saccharomyces boulardii on intestinal mucosa. Dig. Dis. Sci., 51, 1485, 2006. 42. Czerucka, D., Piche, T., and Rampal, P. Review article: yeast as probiotics—Saccharomyces boulardii. Aliment. Pharmacol. Ther., 26, 767, 2007. 43. Dalmasso, G. et al. Saccharomyces boulardii inhibits inflammatory bowel disease by trapping T cells in mesenteric lymph nodes. Gastroenterology, 131, 1812, 2006. 44. van der Aa Kühle, A., Skovgaard, K., and Jespersen, L. In vitro screening of probiotic properties of Saccharomyces cerevisiae var. boulardii and food-borne Saccharomyces cerevisiae strains. Int. J. Food Microbiol., 101, 29, 2005. 45. Sougioultzis, S. et al. Saccharomyces boulardii produces a soluble anti-inflammatory factor that inhibits NF-kappaBmediated IL-8 gene expression. Biochem. Biophys. Res. Commun., 343, 69, 2006.

Saccharomyces 46. Burkhardt, O. et al. Saccharomyces boulardii induced sepsis: successful therapy with voriconazole after treatment failure with fluconazole. Scand J. Infect. Dis., 37, 69, 2005. 47. Opinion of the Scientific Committee on a request from EFSA on the introduction of a Qualified Presumption of Safety (QPS) approach for assessment of selected microorganisms referred to EFSA. EFSA J., 587, 1, 2007. 48. Enache-Angoulvant, A. and Hennequin, C. Invasive Saccharomyces infection: a comprehensive review. Clin. Infect. Dis., 41, 1559, 2005. 49. Almanza, L. et al. Quatre heures pour un record, ou une cellulite vraiment foudroyante: Saccharomyces cerevisiae peut-elle en être la cause? Ann. Fr. Anesfh. Rianim., 1, 130, 1998. 50. Wilson, J.D., Jones, B.M. and Kinghorn, G.R. Bread-making as a source of vaginal infection with Saccharomyces cerevisiae. Report of a case in a woman and apparent transmission to her partner. Sex Transm. Dis., 15, 35, 1988. 51. Nyirjesy, P. et al. Saccharomyces cerevisiae vaginitis: transmission from yeast used in baking. Obstet. Gynecol., 86, 326, 1995. 52. de Llanos, R. et al. Food and probiotic strains from the Saccharomyces cerevisiae species as a possible origin of human systemic infections. Int. J. Food Microbiol., 110, 286, 2006. 53. de Llanos, R., Fernandez-Espinar, M.T. and Querol, A. A comparison of clinical and food Saccharomyces cerevisiae isolates on the basis of potential virulence factors. Antonie Van Leeuwenhoek, 90, 221, 2006. 54. Lherm, T. et al. Seven cases of fungemia with Saccharomyces boulardii in critically ill patients. Intensive Care Med., 28, 797, 2002. 55. Cassone, M. et al. Outbreak of Saccharomyces cerevisiae subtype boulardii fungemia in patients neighboring those treated with a probiotic preparation of the organism. J. Clin. Microbiol., 41, 5340. 2003. 56. Clemons, K.V. et al. Application of DNA typing methods and genetic analysis to epidemiology and taxonomy of Saccharomyces. J. Clin. Microbiol., 35, 1822, 1997. 57. Clemons, K.V. et al. Comparative pathogenesis of clinical and nonclinical isolates of Saccharomyces cerevisiae. J. Infect. Dis., 169, 859, 1994. 58. de Llanos, R. et al. Molecular characterisation of clinical Saccharomyces cerevisiae isolates and their association with non-clinical strains. System. Appl. Microbiol., 27, 427, 2004. 59. Sobel, J.D. et al. Vaginitis due to Saccharomyces cerevisiae: epidemiology, clinical aspects, and therapy. Clin. Infect. Dis., 16, 93, 1993. 60. McCullough, M.J. et al. Epidemiological investigation of vaginal Saccharomyces cerevisiae isolates by a genotypic method. J. Clin. Microbiol., 36, 557, 1998. 61. Kortekangas-Savolainen, O. et al. Immediate hypersensitivity to bakery, brewery and wine products in yeast-sensitive atopic dermatitis patients. Clin. Exp. Allergy, 24, 836, 1994. 62. van Burik, J.H. and Magee, P. Aspects of fungal pathogenesis in humans. Annu. Rev. Microbiol., 55, 743, 2001. 63. McCusker, J.H. et al. Saccharomyces cerevisiae virulence phenotype as determined with CD-1 mice is associated with the ability to grow at 42 °C and form pseudohyphae. Infect. Immun., 62, 5447, 1994. 64. Wheeler, R.T., et al. A Saccharomyces cerevisiae mutant with increased virulence. Proc. Natl. Acad. Sci. USA, 100, 2766, 2003.

635 65. Hodgetts, S. et al. Over-expression of Saccharomyces cerevisiae hsp90 enhances the virulence of this yeast in mice. FEMS Immunol. Med. Microbiol., 16, 229, 1996. 66. Kurtzman, C.P. and Robnett, C.J. Identification and phylogeny of ascomycetous yeasts from analysis of nuclear large subunit (26S) ribosomal DNA partial sequences. Antonie Van Leeuwenhoek, 62, 331, 1998. 67. McCullough, M.J. et al. Intergenic transcribed spacer PCR ribotyping for differentiation of Saccharomyces species and interspecific hybrids. J. Clin. Microbiol., 36, 1035, 1998. 68. Esteve-Zarzoso, B. et al. Identification of yeasts by RFLP analysis of the 5.8S rRNA gene and the two ribosomal internal transcribed spacers. Int. J. Syst. Bacteriol., 49, 329, 1999. 69. Martorell, P., Querol, A., and Fernandez-Espinar, M.T. Rapid identification and enumeration of Saccharomyces cerevisiae cells in wine by real-time PCR. Appl. Environ. Microbiol., 71, 6823, 2005. 70. Chang, H.W. et al. Quantitative real-time PCR assays for the enumeration of Saccharomyces cerevisiae and the Saccharomyces sensu stricto complex in human feces. J. Microbiol. Methods, 71, 191, 2007. 71. Campa, D. et al. A DNA microarray based on Arrayed-Primer Extension (APEX) technique for identification of pathogenic fungi responsible for invasive and superficial mycoses. J. Clin. Microbiol., 46, 909, 2008. 72. Demuyter, C. et al. Predominance of Saccharomyces uvarum during spontaneous alcoholic fermentation, for three consecutive years, in an Alsatian winery. J. Appl. Microbiol., 97, 1140, 2004. 73. Lopez, V. et al. A simplified procedure to analyse mitochondrial DNA from industrial yeasts. Int. J. Food Microbiol., 68, 75, 2001. 74. Schuller, D. et al. Genetic characterization of commercial Saccharomyces cerevisiae isolates recovered from vineyard environments. Yeast, 24, 625, 2007. 75. De Barros Lopes, M. et al. PCR differentiation of commercial yeast strains using intron splice site primers. Appl. Environ. Microbiol., 62, 4514, 1996. 76. Ness, F. et al. Identification of yeast strains using the polymerase chain reaction. J. Sci. Food Agri., 62, 89, 1993. 77. Legras, J.L. and Karst, F. Optimization of interdelta analysis for Saccharomyces cerevisiae strain characterization. FEMS Microbiol. Lett., 221, 249, 2003. 78. Legras, J.L. et al. Selection of hypervariable microsatellite loci for the characterization of Saccharomyces cerevisiae strains. Int. J. Food Microbiol., 102, 73, 2005. 79. Marinangeli, P. et al. Minisatellites in Saccharomyces cerevisiae genes encoding cell wall proteins: a new way towards wine strains characterisation. FEMS Yeast Res., 4, 427, 2004. 80. Schuller, D. et al. Survey of molecular methods for the typing of wine yeast strains. FEMS Microbiol. Lett., 231, 19, 2004. 81. Le Jeune, C. et al. Evolution of the population of Saccharomyces cerevisiae from grape to wine in a spontaneous fermentation. Food Microbiol., 23, 709, 2006. 82. Posteraro, B. et al. Molecular tools for differentiating probiotic and clinical strains of Saccharomyces cerevisiae. Int. J. Food Microbiol., 103, 295, 2005. 83. Vaudano, E. and Garcia-Moruno, E. Discrimination of Saccharomyces cerevisiae wine strains using microsatellite multiplex PCR and band pattern analysis. Food Microbiol., 25, 56, 2008.

636 84. Karakousis, A. et al. An assessment of the efficiency of fungal DNA extraction methods for maximizing the detection of medically important fungi using PCR. J. Microbiol. Methods, 65, 38, 2006. 85. Meroth, C.B. et al. Monitoring the bacterial population dynamics in sourdough fermentation processes using PCRdenaturing gradient gel electrophoresis. Appl. Environ. Microbiol., 69, 475, 2003. 86. Meroth, C.B., Hammes, W.P. and Hertel, C. Identification and population dynamics of yeasts in sourdough fermentation processes by PCR-denaturing gradient gel electrophoresis. Appl. Environ. Microbiol., 69, 7453, 2003. 87. Muñoz, P. et al. Saccharomyces cerevisiae fungemia: an emerging infectious disease. Clin. Infect. Dis., 40, 1625, 2005.

Molecular Detection of Foodborne Pathogens 88. Selvarangan, R. et al. Rapid identification of commonly encountered Candida species directly from blood culture bottles. J. Clin. Microbiol., 41, 5660, 2003. 89. Lorenz, M.C. and Heitman, J. The MEP2 ammonium permease regulates pseudohyphal differentiation in Saccharomyces cerevisiae. EMBO J., 17, 1236, 1998. 90. Lee, K.S. et al. The Saccharomyces cerevisiae PLB1 gene encodes a protein required for lysophospholipase and phospholipase B activity. J. Biol. Chem., 269, 19725, 1994. 91. Edwards-Ingram, L. et al. Genotypic and physiological characterization of Saccharomyces boulardii, the probiotic strain of Saccharomyces cerevisiae. Appl. Environ. Microbiol., 73, 2458, 2007.

Section V Foodborne Protozoa

45 Acanthamoeba Hélène Yera

Université Paris Descartes

Pablo Goldschmidt, Christine Chaumeil

Laboratoire du Centre National d‘Ophtalmologie des Quinze-Vingts

Muriel Cornet

Hôpital Hôtel-Dieu

Marie-Laure Dardé CHU

Contents 45.1 Introduction.................................................................................................................................................................... 639 45.1.1 Taxonomy and Biology..................................................................................................................................... 639 45.1.2 Acanthamoeba keratitis.................................................................................................................................... 640 45.1.2.1 Clinical Characteristics and Pathogenesis ...................................................................................... 640 45.1.2.2 Conventional Diagnosis .................................................................................................................. 640 45.1.2.3 Molecular Diagnosis........................................................................................................................ 641 45.1.3 Other Acanthamoeba infections....................................................................................................................... 641 45.1.3.1 Clinical Characteristics and Pathogenesis....................................................................................... 641 45.1.3.2 Conventional Diagnosis................................................................................................................... 642 45.1.3.3 Molecular Diagnosis ....................................................................................................................... 642 45.2 Methods.......................................................................................................................................................................... 643 45.2.1 Reagents and Equipment................................................................................................................................... 643 45.2.2 Sample Preparation .......................................................................................................................................... 643 45.2.3 Detection Procedures........................................................................................................................................ 644 45.2.3.1 Standard PCR Detection . ............................................................................................................... 644 45.2.3.2 Real-time TaqMan PCR Detection................................................................................................ 644 45.2.3.3 Genotyping Based on Sequencing Analysis.................................................................................... 645 45.3 Conclusions and Future Perspectives............................................................................................................................. 646 References.................................................................................................................................................................................. 647

45.1  Introduction 45.1.1 Taxonomy and Biology Acanthamoeba spp. are found as free-living amoebae in the aquatic environment (bottled water, tap water, chlorinated swimming pools, jacuzzis, seawater, sewage, power plant, etc.), soil and air, and can be isolated from rhino pharynx and stools of healthy humans.1 According to the traditional hierarchical systems of taxonomy, the genus Acanthamoeba belongs to the Family Acanthamoebidae, Subclass Gymnamoebia, Class Lobosoea, Superclass Rhizopoda, Subkingdom Protista.2 Recently, the International Society of Protozoologists adopted a system based on modern morphological approaches,

biochemical pathways and molecular phylogenetic analysis.3 According to this scheme, Acanthamoeba is classified under Super Group Amoebozoa: Acanthamoebidae. During its life cycle, Acanthamoeba undergoes two stages: a vegetative trophozoite and a resistant cyst. Spiny surface projections, called acanthopodia, are presented on the trophozoites and distinguish Acanthamoeba from other free-living amebae that infected human (Balamuthia mandrillaris, Naegleria fowleri, and Sappinia diploidea).4 During the trophozoite stage, Acanthamoeba actively feed on bacteria, algae, yeasts, or small organic particles. This stage is maintained at neutral pH, appropriate temperature (i.e., 30°C) and osmolarity (50–80 mOsmol). Harsh conditions 639

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Molecular Detection of Foodborne Pathogens

(i.e., lack of nutriments, extreme variation in environmental pH, temperatures, and osmolarity) induce the transformation of trophozoites into cyst stage. The cysts possess a double wall and pores called ostioles, which are capable of monitoring environmental changes.2 In addition to being airborne, the cysts are resistant to several biocides, chlorination, and antibiotics, and survive low temperature (0–2°C).4 Upon return to favorable growth conditions, the trophozoite emerges from the cyst by dislodging the operculum which covers the pore and by reverting to a trophic form.5 Acanthamoeba spp. are nonopportunistic or opportunistic pathogens in humans and are responsible for corneal infections in immunocompetent individuals and for brain or disseminate infections in immunocompromised individuals. The host immune status, as well as the material characteristics of contact lens, plays a part in pathogenenesis,6 which is also dependent on strain virulence. Pathogenic isolates exhibit higher binding to host cells and increased cytotoxicity.7 Extracellular proteases are major Acanthamoeba virulence factors, being involved in host tissue breakdown.8,9

45.1.2 Acanthamoeba keratitis 45.1.2.1  Clinical Characteristics and Pathogenesis Acanthamoeba keratitis (AK) is a sight-threatening infection. The greatest populations at risk are soft contact lens wearers associated with poor contact lens hygiene10 or ineffective lens disinfection6 and they have been mainly described in industrialized countries. In developing regions, data concern mostly non-contact lens wearers and subjects with corneal injuries with plants or soil.11,12 AK cases could also be observed after dirty water projection or dust in eye.11,13 Animal studies have shown that contact lenses serve as vectors for transmitting trophozoites to the corneal epithelium and that abrasion or mild trauma to the corneal surface enhances the level of trophozoite binding and is required for the establishment of corneal infection.8 The pathogenesis of AK begins when trophozoites adhere to mannosylated glycoproteins upregulated on the surfaces of wounded corneal epithelium and secrete a mannose-induced cytolytic protein.8,14 A

Trophozoites breech Bowman’s membrane and enter the collagenous stroma. They continue to produce several proteases, including a collagenase and a plasminogen activator, that contribute to the dissolution of the corneal stroma. Then, trophozoites often cluster around corneal nerves, producing radial keratoneuritis and severe pain.8 At the onset of the infection, the clinical diagnosis is difficult to assess due to the lack of distinct pathological signs and to the similarity with herpetic keratitis. Most patients complain of photophobia, pain, tearing, and blush. When the clinical signs become typical (keratitis with a stromal infiltrate) and severe, Acanthamoeba evolves deep in the cornea. AK usually reaches one eye, but it can be occasionally bilateral. Topical treatment requires a combination of eye drops containing biguanides, diamidines and antibiotics (i.e., 0.02% polyhexamethylene biguanide with hexamidine and colimycin 125000 UI/ml) and can be efficient if administered at disease onset (i.e. each hour for 3 days, then each two hours the day and each four hours the night for 8 days, then five to four times a day for a number of weeks according to the microbiological results and the clinical evolution). In severe cases presenting a corneal ulceration and stromal infiltrate, the treatment should be prolonged to avoid recurrences and the corneal scars may require keratoplasty to recover visual acuity. On the above, an early AK diagnosis significantly improves the prognosis if appropriate therapeutic is administrated.15 45.1.2.2  Conventional Diagnosis Conventional laboratory diagnosis of AK has been dependent on direct examination of deep corneal scrapings or corneal biopsies and culture. Direct examination. Corneal samples can be smeared onto glass slides and stained with Gram, ­May–Grünwald–Giemsa pH 7.4, Giemsa, hematoxylin-eosin-safran, lactophenol cotton blue or calcofluor-white for microscopic examination. The Acanthamoeba trophozoites are 15–50 µm in length and have a single nucleus with a prominent central nucleolus. The cysts are 10–25 µm long and have a double wall; most of the species at the points where the two layers joined B

Figure 45.1  Acanthamoeba cysts in corneal biopsy smear stained with hematoxylin-eosin-safran 200 × magnification (A) and 1000 × magnification (B). Chaumeil, C.

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Acanthamoeba

are pore closed by an operculum, giving frequently rise to a polygonal shape (Figure 45.1). Culture. The material obtained from the corneal samples is discharged into a small portion of 2% agar gel and cultivated into flasks containing 5 ml of 0.9% sodium chloride and a killed Escherichia coli suspension (0.5 McFarland). The flasks are incubated at 30°C for 20 days and observed daily for detection of amebae by inverted microscopy. Acanthamoeba trophozoites show slow movements by extending hyaline pseudopodia, and may contain one or more contractile vacuoles. The cysts are also characterized by their double wall. Contact lens case solutions are centrifuged and the pellets are cultivated into flasks as described above. Contact lenses are directly introduced into flasks. Because Acanthamoeba have been detected in lens cases of up to 8% asymptomatic subjects,6 the Acanthamoeba isolation from contact lenses or lens case solutions warrants the analysis of corneal samples if the patient shows symptoms. These conventional methods lack sensitivity. About 60% of AK cases are misdiagnosed using direct detection methods16–18 and positive cultures may be obtained only after prolonged incubation, particularly if antibiotics with potential amoebicidal activity have been used previously.16 45.1.2.3  Molecular Diagnosis Early AK diagnosis and treatment are essential for improving the visual prognosis.15 Several studies have confirmed the usefulness of PCR for the early and sensitive diagnosis of AK in corneal samples and positive PCR results could be obtained in samples from patients previously treated with antibiotics presenting amebicidal activity.16,19,20 DNA extraction methods. Acanthamoeba DNA was purified from corneal scrapings or cultures using the UNSET (urea-NaCl-SDS-EDTA-tris hydrochloride) buffer lysis method21 followed by phenol-chloroform-isoamyl alcohol (25:24:1) and ethanol precipitation.22,23 A modified Chelex24 procedure was used for clinical samples containing lower numbers of amebae.22 Acanthamoeba DNA extraction was performed from corneal samples and cultures by using commercial kits: QIAmp DNA Mini kit (Qiagen, France) by using tissue protocol,19,20 DNA mini prep kit (Biogene)25 and MagNA Pure® Nucleic Acid isolation kit (Roche Diagnostics, France) with the MagNa Pure® Compact (Roche Diagnostics) automate.26 Because the cysts wall is very resistant, the manufacturer protocols have been modified to increase their lysis capacities by adding sodium hydroxide19,20 or proteinase K pre-treatment.26 The pre-treatment with proteinase K before carrying out the commercial procedures increases the rate of positive results. Moreover, the DNA structures were not affected and the PCR inhibitors were eliminated.26 PCR methods. Acanthamoeba PCR targeting regions of the Acanthamoeba nuclear small-subunit rRNA (Rns) gene12,16,17,19,20,22,23 or the Acanthamoeba nuclear ­large-subunit rRNA gene23 have been reported. On Rns gene, the follow­ ing primer pairs have been used: P1GP.2379for–2632rev

primers,16 ACARNA.1383for–1655rev primers,16,17,20 Nelson primers17,19,20 and JDP (JDP1-JDP2) primers.12,20,22,23 Nelson primers were designed to produce an amplimer nearly identical to that of the P1GP primers.17 The combination of primer pairs increased the sensitivity of PCR up to 94%.16,17,20,23 A semi-nested Acanthamoeba PCR was developed with JDP1 and JDP2 as the outer primers in a first PCR then with A1 as the inner forward primer in a second PCR.25 The semi-nested PCR showed better sensitivity than the simplex PCR with JDP1 and JDP2 (100% versus 27%) and 100% specificity. A duplex PCR assay was developed with primers targeting the Acanthamoeba nuclear largesubunit rRNA gene and JDP primers.23 The assay based on the two targets provides more reliable detection than the assay performed only with JDP primers and based on the Rns gene.

45.1.3 Other Acanthamoeba Infections 45.1.3.1  Clinical Characteristics and Pathogenesis Acanthamoeba infections out of the cornea have been rarely described. One case of Acanthamoeba chorioretinitis was reported after penetrating keratoplasties in a patient presenting recurrent keratitis27 and one case of Acanthamoeba endophthalmitis in a patient with acquired immunodeficiency syndrome (AIDS), presenting also a disseminated infection.28 Acanthamoeba causes granulomatous amoebic encephalitis (GAE), primarily in patients with HIV/AIDS or suffering chronic diseases like diabetic or have undergone organ transplantation or are otherwise weakened with no recent history of exposure to recreational freshwater.1,2,4,29,30 A few cases have been documented from individuals with no obvious sings of immunodepression1. Acanthamoeba may disseminate and be responsible of infections of the skin, the lung, the sinus, and the nasal passages, especially in patients with HIV/AIDS.2,4,29,30 Infections of the skin but without dissemination to the central nervous system (CNS) have been observed.1 Because the prodromal period may last several weeks or months, the precise portal of entry is not clearly determined. Trophozoites and/or cysts may enter the body by a nasopharyngeal route or by breaks in the skin, resulting in hematogenous dissemination to the lung and brain. Inhalation of cysts may lead to the same dissemination.1 Trophozoites and cysts are often observed in the brain around blood vessels and within necrotic tissue.29 Serine proteases produced by pathogenic isolates may increase the blood-brain barrier permeability31 and degraded extracellular matrix, fibrinogen, IgG, IgA, albumin, and hemoglobin contributing to brain tissue damage.32 In immunocompromised patients, the chronic inflammatory reaction is minimal and without multinucleated giant cells.1,29 The infections are perhaps either misdiagnosed or undiagnosed due to their nonspecific clinical signs. At the onset, GAE is slow and develops as a chronic disease for several weeks to months. The clinical manifestations include

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headache, stiff neck, confusion, nausea, vomiting, fever, lethargy, cerebellar ataxia, visual disturbance, hemiparesis, and coma.1,4 GAE can mimic meningeal tuberculosis, brain tumors, viral encephalitis, neurocysticercosis, cerebral toxoplasmosis, and acute disseminated encephalomyelitis.30 The prognosis of these infections is poor. The therapeutic strategies require a combination of agents with a reduced successful clinical outcome if treatment is administrated late.1 45.1.3.2  Conventional Diagnosis Histologic examination. Detection of Acanthamoeba is often confirmed post-mortem with histopathologic examination. Trophozoites and cysts can be observed in tissues sections after hematoxylin-eosin, Gomori methenamine silver, periodic acid-Schiff or trichrome staining. Only cysts are stained using Gomori methenamine silver. The amebae are distinguishable from host cells by their prominent central nucleolus and their location in the perivascular areas in brain tissue.1 Acanthamoeba spp. infecting the CNS are not readily found in the cerebrospinal fluid (CSF), although a few cases reported their isolation from the CSF.1 Immunostaining with fluorescent antibodies is useful to detect Acanthamoeba trophozoites from tissue sections.33 Testing for Acanthamoeba antibodies. Detection of circulating antibodies by indirect immunofluorescence can be utilized to screen acanthamoebiasis in patients presenting clinical, laboratory, and neuroimagings data suggestive of amebic encephalitis.34 Low antibody titers are found in more than 50% of asymtomatic individuals.35 However, high titers (1:128 or more) or significant increase in titers have been reported in GAE.36,37 Nevertheless, antibody testing seems not to reflect the real picture of infection in immunocompromised patients.37 Culture. Disrupted tissues or CSF are inoculated on nonnutriment agar plates covered with killed E. coli and incubated at 30°C or 37°C37–39 or on sea salt agar plates seeded with a Phyllobacterium species.40 The plates are observed daily for detection of amebae by inverted microscopy. 45.1.3.3  Molecular Diagnosis Recently, PCR has been employed as an adjunct in the diagnosis of noncorneal acanthamoebiasis.30,41–45 The assays can be performed in CSF, bronchoalveolar lavages, brain tissues, and others tissues. Positive PCR results can also be obtained in formalin-fixed paraffin-embedded samples, but this testing scheme shows limited sensitivity and reproducibility.30 DNA extraction methods. Paraffin-embedded tissues are deparaffined in xylene, washed in 100% ethanol and air dried.42 Acanthamoeba DNA extraction is performed from fluids, unfixed, and deparaffined tissues or cultures by adding lysis buffer 46 followed by precipitation with isopropanol and washing with 70% ethanol.47 It is also performed from deparaffined brain tissue or cultures by using commercial kit: DNeasy tissue kit (Qiagen).38,42

Molecular Detection of Foodborne Pathogens

PCR methods. Acanthamoeba PCR targeting the Acanthamoeba mitochondrial small-subunit rRNA (rns) gene with the Aca16Sf1010-Aca16Sr1180 primers30,41,44 or the Acanthamoeba Rns gene with the JDP primers have been described.42,43,45 A real-time TaqMan PCR assay has been developed for the detection of Acanthamoeba spp. in cultures and clinical samples.48 Another real-time TaqMan PCR assay has been designed for the quantification of Acanthamoeba cells from cultures.49 The both assays targeted the Acanthamoeba Rns gene but the first assay is more broadly reactive than the second when testing isolates from genotypes T1, T4, T7 and T10.48 Recently, a real-time TaqMan PCR assay targeting the Acanthamoeba rns gene was devised for detecting strains available from the American Type Culture Collection (ATCC) and related to genotypes T1, T2, T4, T5, T7 to T9.50 Molecular typing of Acanthamoeba strains. Because Acanthamoeba strains exhibit different pathogenic potentials, it is necessary to characterize them for control and prevention purposes. The genus Acanthamoeba can sometimes be differentiated by its morphology into three morphological groups.51 Classification based on this criterion is not reliable due to alternating cyst appearance and species-level identification requires more complex methods.52 The molecular classification of Acanthamoeba isolates is based on amplification and sequencing of the nearly complete Rns genes.53,54 Fifteen genotypes, designated T1–T15, have been described.53–57 Parts of the Rns gene contain sufficient sequence differences that phylogenetically relevant. Such sequence corresponds to the variable region DF3 of the amplimer ASA.S1, obtained with the JDP primer pair, which discriminates most of the Acanthamoeba genotypes.22 Sequence analysis of complete Rns genes or of ASA.S1 amplimers is useful for the characterization of the Acanthamoeba genotypes in clinical or environmental samples. Around 90% of Acanthamoeba isolates associated with AK in different areas belong to genotype T4.20,22,23,58–62 Other closely related genotypes (T3, T11) and more distantly related genotypes (T2, T6, T5) have been identified in AK.20,22,59,61,63 The genotype T4 is also predominant in non-AK infections and in environmental sources.59,61 The genotype T4 is predominant in GAE, but isolates from genotypes T1, T2, T7, T10, and T12 have been also described.4,45,59 The analysis of the complete Acanthamoeba mitochondrial small-subunit rRNA (rns) genes confirmed the identification of the Acanthamoeba genotypes based on Rns genes.64 Recently, examination of the Acanthamoeba first internal transcribed spacer (ITS1) demonstrates that ITS1 is a more variable region than the Rns gene and that the ITS1 clusters correlate well with the Rns clusters.65 In fact, the variability in ITS1 is ten-fold higher than that in the Rns gene, therefore it can be used to discriminate between different genotypes or even within genotype T4.

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Acanthamoeba

45.2  Methods 45.2.1 Reagents and Equipment Stage

Reagents

Corneal sample collection

Equipment Sterile sponges, stainless vaccinostyles or surgical blades

DNA extraction

Sterile phosphate-buffered saline pH 7.2 (bioMérieux, France) QIAmp DNA Mini kit (Qiagen, France) Ethanol 100% MagNA Pure® Nucleic Acid isolation kit (Roche Diagnostics, France)

Laminar flow cabinet Dry bath incubator Refrigerated centrifuge MagNa Pure® Compact automate (Roche Diagnostics)

Classical PCR

AmpliTaq Gold DNA polymerase (Applied Biosystem, France) for the simplex PCR assays Or Taq DNA Pol (Q-Biogene, MP Biomedicals, France) for the triplex PCR assay Uracil-DNA Glycosylase heat-labile (Roche Diagnostics, France) Each DNTP 25 µmoles (GE Healthcare, France) Seaken GTC Agarose (Tebu, France) Tris/Acetate/EDTA (TAE) buffer (Gibco, France) Ethidium bromure (Eurobio, France) Molecular weight V (Roche Diagnostics)

Laminar flow cabinet Vortex agitator Centrifuge Thermal Cycler (Applied Biosystems) Gel electrophoresis apparatus Ultraviolet lamp

Real-time TaqMan PCR

TaqMan Universal Master Mix containing Uracil-DNA Glycosylase (Applied Biosystem) Or FAST TaqMan Universal Master Mix (Applied Biosystem)

Laminar flow cabinet Vortex agitator Centrifuge ABI Prism 7000 sequence detector system (Applied Biosystems) Or ABI Prism 7500 sequence detector system (Applied Biosystems)

Genotyping

QIAquick PCR Purification kit (Qiagen)

Refrigerated centrifuge Automated DNA sequencer

45.2.2 Sample Preparation Sample collection. In patients with clinical signs suggesting AK, the identification (and if possible semi-quantification) of Acanthamoeba in the corneal structures should be conducted.

Samples consist of either corneal scrapings or corneal biopsies. Scrapings are performed with sterile sponges, stainless vaccinostyles or surgical blades in the periphery of the lesion, deeply in the cornea. Vaccinostyles allow better samplings than sponges. Samples are collected in sterile microtubes and conserved at −20°C before processing in the laboratory. Fluorescein and oxybuprocain solutions, used routinely by ophthalmologists for clinical examination and sampling, inhibit even diluted 20 times the PCR.66 To minimize the risks of misdiagnosis, the eye surface should be rinsed intensively with 0.9% sodium chloride solution before sampling. DNA extraction from corneal samples. For DNA extraction from corneal samples, the samples are discharged in 200 µl sterile phosphate-buffered saline (pH 7.2). DNA is extracted manually by using the commercial kit: QIAmp DNA Mini kit (Qiagen, France) with the tissue protocol.20 A pre-treatment with proteinase K at 56°C for one night before using QIAmp DNA Mini kit shows highest yield of positivity than a pre-treatment with hydroxide sodium then proteinase K for 30 min. Briefly, the sample is mixed with 180 µl of buffer ATL and 20 µl of proteinase K, and incubated for one night at 56°C. Then 200 µl of buffer AL is added and the sample is incubated for 10 min at 70°C. Next, 200 µl of ethanol 100% is added. The mix is applied in QIAamp column and centrifuged for 1 min at 6000 × g (8000 rpm) to bind DNA to the silica membrane inside the column. The DNA is washed first with 500 µl of buffer AW1, which is eliminated by centrifugation 1 min at 6000 × g (8000 rpm) then with 500 µl of buffer AW2, which is eliminated by centrifugation 5 min at 17,900 × g (13,000 rpm). A last centrifugation is performed for 1 min at 17,900 × g (13,000 rpm) to eliminate residual buffer. The DNA is eluted by adding 100 µl of sterile distilled water in the column, incubating 5 min and centrifuging for 1 min at 6000 × g (8000 rpm). DNA can be also extracted automatically by the MagNa Pure® Compact (Roche Diagnostics, France) automate using the MagNA Pure® Nucleic Acid isolation kit (Roche Diagnostics) after 10 min proteinase K pre treatment.26 Briefly, the samples are mixed with 200 µl of ATL buffer and 40 µl of proteinase K, and incubated at 56°C for 10 min. The mix is introduced in the automate for DNA isolation based on magnetic-bead technology. DNA is eluted in 50 µl of sterile distilled water. DNA extraction from noncorneal samples. Acanthamoeba DNA is available for amplification using QIAmp DNA Mini kit (Qiagen)20,26 and MagNA Pure® Nucleic Acid isolation kit (Roche Diagnostics).26 The protocols described below have not been tested in noncorneal samples, but they could be used for the extraction of Acanthamoeba DNA from such samples: 200 µl of biological fluid or 10–25 mg of tissues previously cut up or disrupted. In addition, Acanthamoeba DNA could be isolated from deparaffined brain tissue by using another solid column-based extraction kit, DNeasy tissue kit (Qiagen).38,42

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Molecular Detection of Foodborne Pathogens

45.2.3   Detection Procedures 45.2.3.1 Standard PCR Detection Principle: DNA extracts can be examined with three different simplex PCR assays targeting the Acanthamoeba nuclear small-subunit Rns gene with the Nelson (forward: 5′- GTTTGAGGCAATAACAGGT-3′ and reverse: 5′-GAATTCCTCGTTGAAGAT-3′), ACARNA (ACARNA. 1383for: 5′-TCCCCTAGCAGCTTGTG-3′ and ACARNA. 1655rev: 5′- GTTAAGGTCTCGTTCGTTA-3′) and JDP (JDP1: 5′-GGCCCAGATCGTTTACCGTGAA-3′ and JDP2: 5′-TCTCACAAGCTGCTAGGGGAGTCA-3′) primers. Procedure: (1) Prepare PCR mixture (50 µl) containing 5 µl of GeneAmp 10 × PCR Gold buffer (Applied Biosystem, France), 5 µl of 25 mM of MgCl2, 8 nmol of each dNTP (dATP, dTTP, dCTP, dGTP), 16 nmol of dUTP, 10 pmol of each primer set (i.e., Nelson, ACARNA, or JDP), 0.4 U of UracilDNA Glycosylase heat-labile (Roche Diagnostics, France), 3 U of AmpliTaq Gold DNA polymerase (Applied Biosystems), sterile distilled water to a volume of 40 µl, and 10 µl of DNA (neat or diluted ten-fold). Include a tube with 40 µl of PCR mixture and 10 µl of sterile water as a negative control. (2) Subject the PCR tubes to the following cycling program: one cycle of 94°C for 5 min; 44 cycles of 94°C for 30 sec, 58°C for 1 min, and 72°C for 40 sec; one cycle of 72°C for 15 min. A

(3) Separate PCR products by 2% agarose gel containing ethidium bromide and visualize on ultraviolet lamp. Note: DNA extracts can also be tested with a triplex PCR assay using the three primer sets in a 25 µl reaction volume containing 2.5 µl of incubation buffer T. Pol (Q-Biogene, MP Biomedicals, France), 5 µl of 25 mM of MgCl2, 4 nmol of each dNTP (dATP, dTTP, dCTP, dGTP), 8 nmol of dUTP, 5 pmol of each primer, 0.2 U of Uracil-DNA Glycosylase heatlabile (Roche Diagnostics), 1 U of Taq DNA Pol (Q-Biogene, MP Biomedicals), sterile distilled water to 20 µl, and 5 µl of DNA (neat or diluted ten-fold). A tube with 20 µl of PCR mixture and 5 µl of sterile water is included as a negative control. The tubes are subjected to one cycle of 94°C for 10 min; 44 cycles of 94°C for 30 sec, 58.5°C for 1 min 20 sec, and 72°C for 1 min 20 sec; one cycle of 72°C for 10 min. Then, 20 µl of PCR products is electrophoresed in 2% agarose gel containing ethidium bromide and visualized on ultraviolet lamp. Positive samples give a single 229 bp, 272 bp, and 423–551 bp fragments using the Nelson, ACARNA, and JDP primer pairs, respectively. We have detected one to five cysts from Acanthameba suspensions with either simplex or triplex PCR assays according to the DNA extraction protocol above (Figure 45.2). 45.2.3.2 Real-time TaqMan PCR Detection Principle: DNA extracts can be also examined with two real-time TaqMan PCR assays targeting the Acanthamoeba Rns gene with the Acant primers-probe set (AcantF900: 5′- CCCAGATCGTTTACCGTGAA-3′, B

272 bp

229 bp

1

2

3

4

5

6

7

8

9

C

1

2

3

4

5

6

7

3

4

5

6

7

8

9

D

423 – 551 bp

423 – 551 bp

272 bp 229 bp 1

2

3

4

5

6

7

8

9

1

2

8

9

Figure 45.2  PCR detection of Acanthamoeba. Simplex PCR assays with the Nelson primers (A), the ACARNA primers (B), the JDP primers (C). Triplex PCR assay with the three primer pairs (D). In A–D, lanes 1–4, a pre-treatment with proteinase K overnight before applying QIAmp® DNA Mini kit tissues extraction protocol (Qiagen) (with lane 1 containing 50 cysts; lane 2, 5 cysts; lane 3, 1 cyst; and lane 4, negative control); lane 5, molecular weight marker; lanes 6–9, a pre-treatment with NaOH and then proteinase K for 30 min before applying QIAmp® DNA Mini kit tissues extraction protocol (Qiagen) (with lane 6 containing 50 cysts; lane 7, 5 cysts; lane 8, 1 cyst; lane 9, negative control). bp, base pairs.

Acanthamoeba

AcantR1100: 5′-TAAATATTAATGCCCCCAACTATCC-3′ and probe AcantP1000: 5′-(FAM)-CTGCCACCGAATACATTAGCATGG-(TAMRA)-3′) and the Ac primers-probe set (AcF1: 5′-CGACCAGCGATTAGGAGACG-3′, AcR1: 5′-CCGACGCCAAGGACGAC-3′ and probe AcP1: 5′-(VIC)TGAATACAAAACACCACCATCGGCGC-(TAMRA)3′.48,49 We have adapted the assays for reactions in ABI Prism 7000 sequence detector system (Applied Biosystems). Procedure: (1) Prepare real-time PCR mixture (25 µl) containing 12.5 µl of TaqMan Universal Master Mix containing Uracil-DNA Glycosylase (Applied Biosystems), 20 pmol of each primer-probe set, 5 pmol of TaqMan probe, sterile distilled water to 20 µl, and 5 µl of DNA (neat or diluted ten-fold). Include a tube containing 20 µl of PCR mixture and 5 µl of distilled water as a negative control. (2) Subject the reaction mixture to one cycle of 50°C for 2 min; one cycle of 95°C for 10 min; 55 cycles of 95°C for 15 sec, and 60°C for 1 min; using an ABI Prism 7000 sequence detector system (Applied Biosystems). (3) Detect and analyze data using the ABI Prism 7000 system software (Applied Biosystems). Note: Positive samples result in an increase in fluorescence during the amplification. The kinetic of fluorescence takes the form of a sigmoidal curve when plotted as cycle number versus fluorescence emission. The point at which fluorescence surpasses the background noise, called the threshold cycle, is proportional to starting template concentration. Recently, a real-time TaqMan PCR assay targeting the Acanthamoeba rns gene has been described,50 which employs forward primer: 5′-GCAGTCGCGGTAATACGGA-3′, reverse primer: 5′-ACCACCTACGCACCCTTTACA-3′ and probe: 5′-(FAM)-AGTGTTATTCGCATTGACTGGGTGTAA(TAMRA)3′. The PCR assay is performed in a 25-µl volume with 12.5 µl of FAST TaqMan Universal Master Mix (Applied Biosystems) including 12.5 pmol of each primer and 10 pmol of TaqMan probe, and 12.5 µl of DNA. A tube with 12.5 µl of FAST TaqMan Universal Master Mix and 12.5 µl of sterile water is used as a negative control. The reaction is subjected to one cycle of 95°C for 20 sec, 45 cycles of 95°C for 3 sec, and 60°C for 30 sec using an ABI Prism 7500 sequence detector system (Applied Biosystems). In order to assess the detection rate of PCR assays, the real-time and the standard PCR assays have been tested with Acanthamoeba DNA extracts from 17 cultures of ocular samples and one corneal scraping collected from patients with clinical signs or risk factors suggesting AK. We were able to detect zero one to five cysts using the real-time PCR assay. All 18 Acanthamoeba isolates were positive with the realtime PCR assay (Table 45.1): 15 cultures (genotypes T3, T4, T6, and T11) were positive with the three assays, two cultures (genotypes T11 and T14) were negative with the assay using the Ac primers-probe set49 and one culture (genotype T5) was positive only with the assay targeting the Acanthamoeba

645

rns gene.50 All the Acanthamoeba isolates were positive with the three different standard PCR assays, except the culture related to genotype T5 that gave a negative result with the assay using ACARNA primers. The real-time PCR assay targeting the Acanthamoeba rns gene50 gave the best detection rate in the Acanthamoeba genotypes studied. The specificity of the real-time PCR assay targeting the Acanthamoeba Rns gene was evaluated by testing DNA extracts from parasitic, fungal, bacterial, and viral cultures or positive samples: Hartmannella spp., Toxoplasma ­gondii, Leishmania infantum, Cryptosporidum sp., Taenia solium, Candida albicans, C. glabrata, C. krusei, C. ­dubliniensis, Saccharomyces cerevisiae, Cryptococcus ­neoformans, Trichosporon cutaneum, Trichosporon dermatidis, Fonsecaea dermatidis, Aspergillus fumigatus, A. niger, Fusarium solani, F. oxysporum, Scedosporium apiospermum, Penicillium sp., Enterocytozoon bieneusi, Escherichia coli, Pseudomonas aeruginosa, Streptococcus pneumoniae, S. agalactiae, Enterococcus faecalis, Staphylococcus aureus, S. epidermidis, Neisseria meningitidis, Haemophilus influenza, Chlamydia pneumophila, Herpes simplex virus 1 and 2, cytomegalovirus, and varicella-zoster virus. The DNA extracts from Hartmannella spp. cultures were tested also with the other PCR assays. The real-time PCR assays led to negative results with the microorganisms DNA. In classic PCR, only ACARNA primers led to false-positive results with Hartmannella spp. DNA. 45.2.3.3 Genotyping Based on Sequencing Analysis Principle: The DF3 sequences of the amplimer ASA.S1, obtained with the JDP primer pair, can be used to identify most Acanthamoeba genotypes.22 The amplimers are purified using a PCR purification kit (e.g., QIAquick PCR Purification kit, Qiagen) and sequenced in both directions using an automated DNA sequencer. 30 µl of positive PCR reaction is mixed with five volumes of buffer PBI and applied to the QIAquick column. DNA binds to the silica membrane after centrifugation for 1 min at 17,900 × g (13,000 rpm). DNA is then washed with 750 µl of buffer PE, which is eliminated by centrifugation 1 min at 17,900 × g (13,000 rpm). A last centrifugation is performed for 1 min at 17,900 × g (13,000 rpm) to eliminate residual buffer. The DNA is eluted by adding 50 µl sterile distilled water in the column, incubating 5 min and centrifuging for 1 min at 17,900 × g (13,000 rpm). 60 ng of purified amplimer are mixed with 4 pmol of JDP1 primer or JDP2 primer and are sequenced in an automatic sequencing platform. The amplimers producing multiple sequencing ladders are cloned into a vector (e.g., pGEM-T Easy, Promega, France) and amplified in competent cells (JM109 High Efficiency Competent cells, Promega). Preparations of plasmid are performed using a miniprep kit (e.g., QIAprep Spin Miniprep kit, Qiagen). Six clones of each amplimer are sequenced on both strands. Amplimers obtained with the ACARNA primer pair are also sequenced when the sample is negative with the JDP primer pair. 30 ng of purified amplimer are mixed with 4 pmol of

646

Molecular Detection of Foodborne Pathogens

Table 45.1 Molecular Detection of Acanthamoeba Isolated from Clinical Specimens Isolate

Sample

Real-time PCR Ac/Acant/rnsa

PCR Nelson/ACARNA/JDPb

Acanthamoeba genotypec (GenBank accession number)

T3  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4  + / + / +   + / + / +  T4 + T4 (EU146068 + EU146069)  + / + / +   + / + / +  T4 (DQ087306)  + / + / +   + / + / +  T4 (EU146073)  + / + / +   + / + / +  T5 (EU146072)  + /−/ +  −/−/ +  T6 (DQ087289)  + / + / +   + / + / +  T11 (DQ087292)  + / + / +   + / + / +  T11 −/ + / +   + / + / +  T14 −/ + / +  nd/nd/ +  Hartmannella sp −/−/− −/ + /– Hartmannella sp (DQ087330) −/−/− −/ + /− Notes: CL: contact lens, CLC: contact lens case solution, CS: corneal scraping. a Ac, primers and probe designed by Rivière, D. et al., J. Microbiol. Methods., 64, 78, 2006; Acant, primers and probe designed by Qvarnstrom, Y. et al. J. Clin. Microbiology., 44, 3589, 2006; rns, primers and probe designed by Chaumeil, C. et al., Invest. Ophthalmol. Vis. Sci., 2492, A18, 2008. b Nelson, Nelson primers; ACARNA, ACARNA.1383for-1655rev primers; JDP, JDP1- JDP2 primers. c Sequence analysis using BLAST searches. d Isolates shared with Yera, H. et al., Br. J. Ophthalmol., 92, 1139, 2008. e Isolate shared with Yera, H. et al., Eur. J. Clin. Microbiol. Inf. Dis., 26, 221, 2007. * DNA was extracted directly from corneal scraping. BOUJ CHAM PETI TROL BARS SPAE PEIG TORO DIPL BLON S32d S11d S36d S35d S7d 143GUILe AFRI ARO SKAL S8d

CL CL CS CL CLC CS CS CL CL CLC CS CL + CLC CLC CLC CL + CLC CS* CS CLC CL + CLC CLC

ACARNA.1383 for or ACARNA.1655rev and are sequenced in an automatic sequencing platform. The sequences are aligned and compared with GenBank reference sequences using BLAST searches.

45.3 Conclusions and Future Perspectives Early and sensitive diagnosis and on time treatment may prevent the poor outcome of Acanthamoeba infections. The PCR assays described for Acanthamoeba diagnosis mainly target Acanthamoeba Rns gene.16,17,19,20,22,23,48,49 The JDP primers are genus-specific and allow sequencing of the amplimers for further identification of most Acanthamoeba genotypes.22 The combination of several primer pairs increased the sensitivity of PCR.16,17,20 False-positive results were observed with Hartmannella isolates when using the ACARNA primer pair in this study and elsewhere.62 ACARNA and P1GP primers were

predicted to produce amplimer with amoebae from the genera Hartmannella and Balamuthia.22 A false-positive result was described by using the Nelson primers on one corneal scraping from a patient without AK; sequence analysis of the amplimer revealed DNA from Vexilliferidae, another freeliving ameba.20 False-negative results are possible despite the combination of several primer pairs.20 In fact, the quality of corneal scrapings may lead to reduce number of Acanthamoeba in the samples. Additionally, the levels of DNA extracted from clinical samples containing only mature cysts can determine the quality of PCR results.22 PCR inhibitors, including topical solutions, could also cause false-negative results. Therefore, an internal control should be added in assays performed on clinical samples. The use of commercial kits (QIAmp DNA Mini kit, Qiagen, and MagNA Pure® Nucleic Acid isolation kit, Roche Diagnostics) improve partially the sensitivity of the PCR assay by eliminating the inhibitors present in the proteolytic mixtures during the DNA extraction.26

647

Acanthamoeba

The presence of inhibitors was assessed by the delays in the signals amplification of an internal control, suggesting that the negative results reflected the absence of available parasite DNA in the PCR reaction tubes. Real-time PCR are sensitive tools for the detection and quantification of microorganisms, allowing faster analysis. The risk of amplicon dispersion and cross-contamination is minimized. TaqMan PCR assays have recently been described for the detection and quantification of Acanthamoeba from cultures49,50 or artificial and clinical samples.48 We confirmed that the Acant primers-probe set48 is able to detect a larger spectrum of genotypes than the Ac one49 (isolates from genotypes T1, T3, T4, T6, T7, T10, T11, and T14. However, these two assays were unable to detect one isolate related to genotype T5 and obtained from a patient’s contact lens case solution.62 The genotype T5 has mainly been described in the environment61 but one isolate has been observed in a keratitis case63 and another in a disseminated infection.39 The real-time PCR assay targeting the Acanthamoeba rns gene50 gave the best detection rate in the Acanthamoeba genotypes studied here. The pre-treatment with proteinase K before DNA extraction procedures and the combination of primer pairs increase the sensitivity of PCR. The choice of DNA extraction methods has an impact on PCR results.26 The JDP primers appear as the most appropriate for targeting genus-specific sequences, allowing further for identification of the genotypes. The primers-probe set designed for real-time PCR and targeting the Acanthamoeba rns gene50 seems to detect the larger spectrum of genotypes among the different real-time PCR assays. Other methods could be combined with PCR to increase the sensitivity. Some authors recommended combining PCR with smear examination for the rapid diagnosis of AK.12 Other authors used PCR to confirm confocal microscopy diagnosis of AK in contact lens wearers.17 Treatment of Acanthamoeba infections requires a long lasting regime based on the combination of different molecules and with limited successful outcome when late.1 AK and other Acanthamoeba infections can be misdiagnosed particularly at the onset of the disease due to a lack of specific clinical signs.16,17,30 Direct examination presented low sensitivity and requires well-trained microscopists. Culture lacks sensitivity in particular if the patients have been treated by antibiotics with potential amebicidal activity. The usefulness of PCR has been demonstrated for the confirmation of AK and is promising for the diagnosis of GAE or others Acanthamoeba infections. The PCR can be performed on corneal samples, biological fluids, formalin-fixed ­paraffin-embedded or unfixed tissues. Because the genome of Acanthamoeba is highly variable between the isolates, the combination of primer sets is recommended to increase the detection rates of PCR. Finally, a better and wider knowledge of the acanthamoebiasis will lead to an early diagnosis and on time treatment, improving the prognosis.

References











1. Visvesvara, G.S., Moura, H. and Schuster, F.L. Pathogenic and opportunistic free-living amoebae: Acanthamoeba spp., Balamuthia mandrillaris, Naegleria fowleri, and Sappinia diploidea. FEMS Immunol. Med. Microbiol., 50, 1, 2007. 2. Khan, N.A. Acanthamoeba: biology and increasing importance in human health. FEMS Microbiol. Rev., 30, 564, 2006. 3. Adl, S.M. et al. The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. J. Eukaryot. Microbiol., 52, 399, 2005. 4. Marciano-Cabral, F. and Cabral, G. Acanthamoeba spp. as agents of disease in humans. Clin. Microbiol. Rev., 16, 273, 2003. 5. Page, F.C. Re-definition of the genus Acanthamoeba with descriptions of three species. J. Protozool., 14, 709, 1967. 6. Illingworth, C.D. and Cook, S.D. Acanthamoeba keratitis. Surv. Ophthalmol., 42, 493, 1998. 7. Alsam, S. et al. Acanthamoeba interactions with human brain microvascular endothelial cells. Microb. Pathog., 35, 235, 2003. 8. Clarke, D.W. and Niederkorn, J.Y. The pathophysiology of Acanthamoeba keratitis. Trends Parasitol., 22, 175, 2006. 9. Kim, W.T. et al. Comparison of specific activity and cytopathic effects of purified 33 kDa serine proteinase from Acanthamoeba strains with different degree of virulence. Korean J. Parasitol., 44, 321, 2006. 10. Radford, C.F. et al. Risk factors for Acanthamoeba keratitis in Contact lens users: a case-control study. Br. Med. J., 310, 1567, 1995. 11. Sharma, S., Garg, P. and Rao G.N. Patient characteristics, diagnosis, and treatment of non-contact lens related Acanthamoeba keratitis. Br. J. Ophthalmol., 84, 1103, 2000. 12. Pasricha, G. et al. Use of 18S rRNA gene-based PCR assay for diagnosis of Acanthamoeba keratitis in non-contact lens wearers in India. J. Clin. Microbiol., 41, 3206, 2003. 13. Syam, P.P., Narendran, R. and van der Hoek, J. Persistent Acanthamoeba keratitis in a non-contact lens wearer following exposure to bird seed dust. Br. J. Ophthalmol., 89, 388, 2005. 14. Hurt, M. et al. Pathogenic Acanthamoeba spp secrete a ­mannose-induced cytolytic protein that correlates with the ability to cause disease. Infect. Immun., 71, 6243, 2003. 15. Cardine, S. et al. Prise en charge clinique et pronostic des kératites amibiennes. Etude rétrospective à propos de 25 cas. J. Fr. Ophtalmol., 25, 1007, 2002. 16. Lehmann, O.J. et al. Polymerase chain reaction analysis of corneal epithelial and tear samples in the diagnosis of Acanthamoeba keratitis. Invest. Ophthalmol. Vis. Sci., 39, 1261, 1998. 17. Mathers, W.D. et al. Confirmation of confocal microscopy diagnosis of Acanthamoeba keratitis using polymerase chain reaction analysis. Arch. Ophthalmol., 118, 178, 2000. 18. Stothard, D.R. et al. Fluorescent oligonucleotide probes for clinical and environmental detection of Acanthamoeba and the T4 18S rRNA gene sequence type. J. Clin. Microbiol., 37, 2687, 1999. 19. Zamfir, O. et al. Diagnostic par PCR des kératites à Acanthamoeba spp. J. Fr. Ophtalmol., 29, 1034, 2006. 20. Yera, H. et al. Comparison of PCR, microscopic examination and culture in the early diagnosis and characterization of Acanthamoeba isolates from ocular infections. Eur. J. Clin. Microbiol. Infect. Dis., 26, 221, 2007.

648 21. Hugo, E.R. et al. Purification of amoeba mtDNA using the UNSET procedure. In: Soldo, A.T. and Lee, J.J. (eds), Protocols in Protozoology. Lawrence KS: Allen Press, 1992. 22. Schroeder, J.M. et al. Use of subgenic 18S ribosomal DNA PCR and sequencing for genus and genotype identification of Acanthamoeba from humans with keratitis and from sewage sludge. J. Clin. Microbiol., 39, 1903, 2001. 23. Sharma, S. et al. Acanthamoeba keratitis in non-contact lens wearers in India: DNA typing-based validation and a simple detection assay. Arch. Ophthalmol., 122, 1430, 2004. 24. Walsh, P.S., Metzger, D.A. and Higuchi, R. Chelex 100 as a medium for simple extraction of DNA for PCR-based typing from forensic material. Biotechniques, 10, 506, 1991. 25. Dhivya, S. et al. Comparison of a novel semi-nested polymerase chain reaction (PCR) with a uniplex PCR for the detection of Acanthamoeba genome in corneal scrapings. Parasitol. Res., 100, 1303, 2007. 26. Goldschmidt, P. et al. Resistance of Acanthamoeba to classic DNA extraction methods used for the diagnosis of corneal infections. Br. J. Ophthalmol., 92, 112, 2008. 27. Moshari, A. et al. Chorioretinitis after keratitis caused by Acanthamoeba: case report and review of the literature. Ophthalmology, 108, 2232, 2001. 28. Heffler, K.F. et al. Acanthamoeba endophthalmitis in acquired immunodeficiency syndrome. Am. J. Ophthalmol., 122, 584, 1996. 29. Martinez, A.J., and Visvesvara, G.S. Free-living, amphizoic and opportunistic amebas. Brain Pathol., 7, 583, 1997. 30. Yagi, S., Schuster, F.L. and Visvesvara, G.S. Demonstration of Balamuthia and Acanthamoeba mitochondrial DNA in sectioned archival brain and other tissues by polymerase chain reaction. Parasitol. Res., 102, 491, 2008. 31. Alsam, S. et al. Extracellular proteases of Acanthamoeba castellanii (encephalitis isolate belonging to T1 genotype) contribute to increased permeability in an in vitro model of the human blood-brain barrier. J. Infect., 51, 150, 2005. 32. Kong, H.H., Kim, T.H., and Chung, D.I. Purification and characterization of a secretory serine proteinase of Acanthamoeba healyi isolated from GAE. J. Parasitol., 86, 12, 2000. 33. Epstein, R.J. et al. Rapid diagnosis of Acanthamoeba keratitis from corneal scrapings using indirect fluorescent antibody staining. Arch. Ophthalmol., 104, 1318, 1986. 34. Schuster, F.L. et al. Detection of antibodies against free-living amoebae Balamuthia mandrillaris and Acanthamoeba species in a population of patients with encephalitis. Clin. Infect. Dis., 42, 1260, 1986. 35. Chappell, C.L. et al. Standardized method of measuring Acanthamoeba antibodies in sera from healthy human subjects. Clin. Diagn. Lab. Immunol., 8, 724, 2001. 36. Grunnet, M.L., Cannon, G.H., and Kushner, J.P. Fulminant amebic meningoencephalitis due to Acanthamoeba. Neurology., 31, 174, 1981. 37. Bloch, K.C., and Schuster, F.L. Inability to make a premortem diagnosis of Acanthamoeba species infection in a patient with fatal granulomatous amebic encephalitis. J. Clin. Microbiol., 43, 3003, 2005. 38. McKellar, M.S. et al. Fatal granulomatous Acanthamoeba encephalitis mimicking a stroke, diagnosed by correlation of results of sequential magnetic resonance imaging, biopsy, in vitro culture, immunofluorescence analysis, and molecular analysis. J. Clin. Microbiol., 44, 4265, 2006.

Molecular Detection of Foodborne Pathogens 39. Barete, S. et al. Fatal disseminated Acanthamoeba lenticulata infection in a heart transplant patient. Emerg. Infect. Dis., 13, 736, 2007. 40. Petry, F. et al. Early diagnosis of Acanthamoeba infection ­during routine cytological examination of cerebrospinal fluid. J. Clin. Microbiol., 44, 1903, 2006. 41. Shirwadkar, C.G. et al. Acanthamoeba encephalitis in patient with systemic lupus, India. Emerg. Infect. Dis., 12, 84, 2006. 42. MacLean, R.C. et al. Identification of Acanthamoeba sp. in paraffin-embedded CNS tissue from an HIV + individual by PCR. Diagn. Microbiol. Infect. Dis., 57, 289, 2007. 43. Meersseman, W. et al. Rapidly fatal Acanthamoeba encephalitis and treatment of cryoglobulinemia. Emerg. Infect. Dis., 13, 469, 2007. 44. Yagi, S., Schuster, F.L., and Bloch, K. Demonstration of presence of Acanthamoeba mitochondrial DNA in brain tissue and cerebrospinal fluid by PCR in samples from a patient who died of granulomatous amebic encephalitis. J. Clin. Microbiol., 45, 2090, 2007. 45. Walochnik, J. et al. Granulomatous amoebic encephalitis caused by Acanthamoeba amoebae of genotype T2 in a human immunodeficiency virus-negative patient. J. Clin. Microbiol., 46, 338, 2008. 46. Casas, I. et al. New method for the extraction of viral RNA and DNA from cerebrospinal fluid for use in the polymerase chain reaction assay. J. Virol. Methods., 53, 25, 1995. 47. Yagi, S. et al. Detection of Balamuthia mitochondrial 16S rRNA gene DNA in clinical specimens by PCR. J. Clin. Microbiol., 43, 3192, 2005. 48. Qvarnstrom, Y. et al. Multiplex real-time PCR assay for the simultaneous detection of Acanthamoeba spp., Balamuthia mandrillaris, and Naegleria fowleri. J. Clin. Microbiology., 44, 3589, 2006. 49. Rivière, D. et al. Development of a real-time PCR assay for quantification of Acanthamoeba trophozoites and cysts. J. Microbiol. Methods., 64, 78, 2006. 50. Chaumeil, C. et al. A new method to detect in one reaction simultaneously 10 different strains of Acanthamoeba infecting humans. Invest. Ophthalmol. Vis. Sci., 2492, A18, 2008. 51. Pussard, M., and Pons. R. Morphologie de la paroi kystique et taxonomie du genre Acanthamoeba (Protozoa, Amoebida). Protistologica, 13, 557, 1977. 52. Visvesvara, G.S. Classification of Acanthamoeba. Rev. Infect. Dis., 13(Suppl 5), S369, 1991. 53. Gast, R.J. et al. Subgenus systematics of Acanthamoeba: four nuclear 18S rDNA sequence types. J. Eukaryot. Microbiol., 43, 498, 1996. 54. Stothard, D.R. et al. The evolutionary history of the genus Acanthamoeba and the identification of eight new 18S rRNA gene sequence types. J. Eukaryot. Microbiol., 45, 45, 1998. 55. Hewett, M.K. et al. Identification of a new Acanthamoeba jacobsi Sawyer, Nerad and Visvesvara, 1992 (Lobosea: Acanthamoebidae). Acta Protozool., 42, 325, 2003. 56. Horn, M. et al. Novel bacterial endosymbionts of Acanthamoeba spp. related to the Paramecium caudatum symbiont Caedibacter caryophilus. Environ. Microbiol., 1, 357, 1999. 57. Gast, R.J. Development of an Acanthamoeba-specific reverse dot-blot and the discovery of a New ribotype. J. Eukaryot. Microbiol., 48, 609, 2001.

Acanthamoeba 58. Booton, G.C. et al. 18S ribosomal DNA typing and tracking of Acanthamoeba species isolates from corneal scrape specimens, contact lenses, lens cases, and home water supplies of Acanthamoeba keratitis patients in Hong Kong. J. Clin. Microbiol., 40, 1621, 2002. 59. Fuerst, P.A. et al. Genotypic identification of no-keratitis infections caused by the oppotunistically pathogenic ameba genus Acanthamoeba. J. Eukaryot. Microbiol., 50 (Suppl), 512, 2003. 60. Zhang, Y. et al. Identification of 18S ribosomal DNA genotype of Acanthamoeba from patients with keratitis in North China. Invest. Ophthalmol. Vis. Sci., 45, 1904, 2004. 61. Booton, G.C. et al. Identification and distribution of Acanthamoeba species genotypes associated with nonkeratitis infections. J. Clin. Microbiol., 43, 1689, 2005.

649 62. Yera, H. et al. The genotypic characterization of Acanthamoeba isolates from human ocular samples. Br. J. Ophthalmol., 92, 1139, 2008. 63. Spanakos, G. et al. Genotyping of pathogenic Acanthamoebae isolated from clinical samples in Greece—report of a clinical isolate presenting T5 genotype. Parasitol. Int., 55, 147, 2006. 64. Ledee, D.R. et al. Advantages of using mitochondrial 16S rDNA sequences to classify clinical isolates of Acanthamoeba. Invest. Ophthalmol. Vis. Sci., 44, 1142, 2003. 65. Köhsler, M. et al. ITS1 sequence variabilities correlate with 18S rDNA sequence types in the genus Acanthamoeba (Protozoa: Amoebozoa). Parasitol. Res., 98, 86, 2006. 66. Goldschmidt, P. et al. Effects of topical anaesthetics and fluorescein on the real-time PCR used for the diagnosis of Herpesviruses and Acanthamoeba keratitis. Br. J. Ophthalmol., 90, 1354, 2006.

46 Cryptosporidium Una Ryan

Murdoch University

Simone M. Cacciò

Istituto Superiore di Sanità

Contents 46.1 Introduction.................................................................................................................................................................... 651 46.1.1 Classification of Cryptosporidium.................................................................................................................... 651 46.1.2 Biology, Pathogenesis, and Medical Importance.............................................................................................. 652 46.1.3 Sources of Foodborne Cryptosporidiosis......................................................................................................... 652 46.1.4 Methods for Diagnosis of Foodborne Cryptosporidium ................................................................................. 655 46.1.4.1 Conventional Methods for Detection of Cryptosporidium.............................................................. 655 46.1.4.2 Molecular Methods for Detection of Cryptosporidium................................................................... 655 46.1.4.3 Detection of Cryptosporidium in Fruit and Vegetables................................................................... 657 46.1.4.4 Detection of Cryptosporidium in Meat, Fish, and Shellfish............................................................ 658 46.1.4.5 Detection of Cryptosporidium in Drinks........................................................................................ 658 46.2 Methods.......................................................................................................................................................................... 659 46.2.1 Reagents and Equipment ................................................................................................................................. 659 46.2.2 Sample Preparation........................................................................................................................................... 659 46.2.3 Detection Procedures........................................................................................................................................ 660 46.2.3.1 Nested PCR Targeting a 825-bp 18S rDNA Fragment ................................................................... 660 46.2.3.2 Nested PCR Targeting a 587-bp 18S rDNA Fragment ................................................................... 660 46.2.3.3 Nested PCR Targeting the GP60 Locus ......................................................................................... 661 46.3 Conclusion and Future Perspectives............................................................................................................................... 661 References.................................................................................................................................................................................. 661

46.1  Introduction

46.1.1 Classification of Cryptosporidium

Cryptosporidium is an enteric parasite, which has a global impact on the health, survival, and economic development of millions of people and animals worldwide for which there is currently no effective chemotherapy. The oocysts produced by Cryptosporidium are extremely hardy and are easily transmitted through contaminated water sources and fresh produce. The rising number of documented cases of cryptosporidiosis, and the potential public health and economic consequences of outbreaks have highlighted the need for rapid, sensitive, and reliable protocols for the detection and differentiation of Cryptosporidium spp. in complex matrices.1 Molecular tools have long been employed in the detection of Cryptosporidium in feces and water but however, relatively few PCR-based methods have been adapted to such complex matrices as foods and beverages.2–4 The diversity of food types, their heterogeneous nature, and the presence of PCR inhibitory substances that are carried over during DNA template preparation all contribute to the difficulty of detecting low levels of Cryptosporidium oocysts in foods and drinks.4

The taxonomic status of Cryptosporidium has been the subject of some debate. Cryptosporidium is a protozoan parasite belonging to the phylum Apicomplexa and until recently, Cryptosporidium was classified as a coccidian, based on life cycle features.5 However, new molecular and biological evidence suggests that it is more closely related to gregarine parasites.6–10 Species within the genus Cryptosporidium have previously been identified based on morphological features and host specificity. Oocysts of Cryptosporidium however are small in size (4–6 µm) and virtually identical between many species and the parasite is not strictly host specific. As a result of this, genetic characterization has become an essential element in defining new species and validating established species. Currently 20 species of Cryptosporidium are regarded as valid (Table 46.1) and over 40 genotypes are recognized, with new genotypes continually being identified.11–16 Five Cryptosporidium species/genotypes are commonly identified in humans; C. hominis, C. parvum, C. meleagridis, C. 651

652

Molecular Detection of Foodborne Pathogens

Table 46.1 Currently Recognized Species of Cryptosporidium Species

Major Host

C. hominis

Humans

C. parvum

Humans, livestock Birds, humans Red kangaroo Grey Kangaroo Cattle Cattle Pigs Cats Dogs Guinea pigs Poultry

C. meleagridis C. fayeri C. marcopodum C. bovis C. ryanae C. suis C. felis C. canis C. wrairi C. baileyi C. galli

Finches, chickens

C. muris C. andersoni C. fragile C. serpentis C. varanii

Rodents Cattle Frogs Snakes, lizards Lizards

C. molnari C. scophthalmi

Fish Fish

Minor Host Dugongs, sheep, cattle

Site of Infection Small intestine Small intestine

Sheep Humans Humans Humans Wide variety of bird species Capercailles, Pine grosbeaks and others. Sheep

Snakes

Small intestine Unknown Unknown Small intestine Small intestine Small intestine Small intestine Small intestine Small intestine Bursa Proventriculus

Stomach Abomasum Stomach Stomach Stomach and small intestine Stomach Intestine

felis, and C. canis, with C. hominis and C. parvum responsible for the majority of infections,13 although, in some endemic areas C. meleagridis infection rate is as high as C. parvum.17

46.1.2 Biology, Pathogenesis, and Medical Importance Upon ingestion by a suitable host, the sporulated oocysts, which contain four sporozoites, excyst and undergo a complex monoxenous life cycle where all stages of development (asexual and sexual) occur within one host. Following fertilization, oocysts may differentiate into thin walled oocysts, which are involved in autoinfection, or thick walled oocysts which exit the host via the feces permitting further transmission via contaminated food/water/fomites and direct contact with infected animals/humans or contaminated surfaces. Thick walled oocysts are environmentally robust and are highly resistant to environmental stresses, including chlorine treatment of community water supplies. Oocysts of some species such as Cryptosporidium parvum can remain infectious in cool, wet conditions for six months or longer.11

Cryptosporidium can involve either children or adults but is a major cause of diarrhoea in children (<5-years-old) in both developed and developing countries.18,19 There is wide variability in the clinical presentation of infection, which primarily occurs in the small intestine. Patients may be asymptomatic or experience acute or chronic diarrhea depending on their age and immune status.11,18,19 In immunocompetent humans, infection usually results in acute self-limiting diarrhea. In immunocompromised individuals, such as individuals on immunosuppressive drugs, AIDS, and those with concurrent infections such as measles or chickenpox, clinical symptoms mimic those of an immunocompetent individual with greater severity and duration and may progress to a chronic state leading to electrolyte imbalance, wasting and eventual death.19 Infection rarely resolves spontaneously. Studies also indicate these individuals are more susceptible to infection from a wider range of Cryptosporidium species and genotypes. The exact mechanism by which Cryptosporidium causes disease is still unknown,20 despite immunological and molecular methodologies identifying a number of candidate molecules involved in adhesion, protein degradation and modulation of host responses.21 The nature and intensity of the diarrhea suggests enterotoxin involvement but none has been demonstrated. The attachment of a sporozoite to the apical surface of epithelial cells triggers a cascade of secondary signal pathways within the host cell, thereby altering its function. The activation of two systems, the nuclear factor-κB (NF-κB) system results in the release of cytokines and chemokines which induce inflammation and anti-apoptotic survival of directly infected cells, while activation of the c-src system causes dysfunction and the apoptosis of healthy adjacent cells.20,21 The outcome is villus atrophy resulting in malabsorption. Recently, it has been shown that the apoptotic response of infected intestinal epithelial cells is actively suppressed by C. parvum via up-regulation of a host antiapoptotic protein (survivin), favoring parasite infection.22

46.1.3 Sources of Foodborne Cryptosporidiosis Cryptosporidium oocysts have been isolated from several foodstuffs (Table 46.2), particularly fruits, vegetables, and shellfish. The contamination with these produce with oocysts is particularly important from a public health viewpoint, as these products are frequently consumed raw without any thermal processing to inactivate contaminating oocysts.23 Cryptosporidium is a common parasite of humans, and foodborne outbreaks of cryptosporidiosis have been reported worldwide (Table 46.3). The main sources of food contamination comes from (i) consumption of raw fruit, vegetables, and shellfish; (ii) processing of meat, beverages and other foodstuffs; (iii) food handlers and (iv) contamination via insects and birds. Raw fruit and vegetables can become contaminated with Cryptosporidium oocysts via water and manure. Water is an important vehicle for the transmission of Cryptosporidium,

653

Cryptosporidium

Table 46.2 Foodstuffs in which Cryptosporidium Oocysts have been Isolated Food Type

Country

Items

References 24 25 26 27 28 29 30 31 32

Italy Eastern United States and Canada Spain

Cilantro, lettuce Radish, carrot, tomato, cucumber Lettuce, mint, cilantro, parsley Lettuce, mung bean sprouts Seed sprouts Lettuce, parsley, cilantro, blackberries Lettuce Water spinach Dosina exoleta, Veneurupis pullastra, V. rhomboids, Venus verrucosa Dosina exoleta, Veneurupis pullastra, V. rhomboids, Venus verrucosa Dosina exoleta, Ruditapes philippinarum, Veneurupis pullastra, V. rhomboids, Venus verrucosa Chamelia gallina clams Mytilus galloprovincialis

Northern Ireland Spain and EU countries Canada United States Ireland Japan United States Eastern US and Canada Spain Spain and EU countries Netherlands Spain Spain and EU countries Europe UK Ohio

Mytilus edulis Mytilus galloprovincialis Zebra mussel (Dreissena ploymorpha) Bent mussel (Ischadium recurvum) Mytilus edulis Corbicula japonica Crassostrea virginica Crassostrea virginica Ostrea edulis Ostrea edulis Crassostrea gigas Cerastoderma edule Cerastoderma edule Small ruminants including sheep and goats Tripe Apple cider

Vegetables

Costa Rica

Shellfish

Peru Norway Norway Costa Rica Northern Ireland Cambodia Spain and Italy Spain Spain and EU countries

Mussels

Oysters

Cockles Meat/meat products Tripe Apple cider

33 34 35 35 32, 33, 34, 37 38 34 39 40 41 42 43–45 36 33, 46 34 47 37 34 48 49 50

Source: Adapted from Millar, C.B. et al., Trends Food Sci. Technol., 13, 168, 2002 and Ortega, I. and Cama V. In Cryptosporidium and Cryptosporidiosis, Fayer, R and Xiao, L. (Eds.), CRC Press, FL, 2008.

and 50.8% of all water-associated outbreaks of parasitic protozoan disease reported in North American and Europe are caused by Cryptosporidium.62 Contamination of water via agricultural runoff or sewage and subsequent irrigation of crops with untreated irrigation water is a significant cause of contamination of fruit and vegetables. In Central America, 36% of irrigation water contained Cryptosporidium oocysts.63 A study of oocyst transport and subsurface irrigation methods demonstrated the presence of oocysts in soils at different depths.64 The use of water sprinklers at retail outlets can also be a source of contamination. This is further supported by a recent study which reported that the C. parvum oocysts tended to associate

with suspended particles in water and that this promoted the oocyst deposition.65 Fertilization of fruit and vegetables, with manure from cattle and sheep containing viable oocysts of Cryptosporidium is another source of contamination and represents a significant public health risk. It has been demonstrated that it took between eight and 31 days to achieve a 1-log reduction in C. parvum levels in farmyard wastes inoculated with Cryptosporidium and spread on a grass pasture.66 The authors reported that protozoans were significantly more persistent than bacteria and that oocyst recovery was more efficient from wastes with a lower dry matter content.66 With a growing consumer trend away from intensively produced crops toward

654

Molecular Detection of Foodborne Pathogens

Table 46.3 Foodborne Outbreaks of Cryptosporidiosis Implicated/ Suspected Food

No. of Cases

Apple cider

154

Apple cider

31

Apple cider Chicken salad Cow’s milk Cow’s milk Cow’s milk Frozen tripe Fruit/vegetables Green onions Goats milk Unknown Unknown

23 15 50 8 22 1 148 54 2 24 88

Country Maine, USA Connecticut and New York, USA Ohio, USA Minnesota, USA UK Queensland, Australia Mexico UK Washington, DC Washington, DC Australia Wisconsin, USA Washington, DC

Suspected Mode of Transmission

References

Cider made from dropped apples on ground grazed by livestock Washing apples with contaminated water

52

Once and twice-ozonated cider Food handler who also worked at day care centre Faulty on-farm pasteurization unit Consumption of unpasteurized milk Obtained from street vendor Oocysts detected in tripe Foodhandler Unwashed onions Consumption of unpasteurized goat’s milk Company and private home

50 54 55 56 57 49 58 59 60 61 61

53

Source: Adapted from Millar, C.B. et al., Trends Food Sci. Technol., 13, 168, 2002.

organic produce, this could increase the likelihood of contamination and the threat to public health.30 The practice of fertilizing horticultural crops with untreated human sewage in some countries is also a considerable risk for the spread of Cryptosporidium to humans who ingest the produce.67,68 Cryptosporidium oocysts have also been recovered from a variety of shellfish species (Table 46.2). Shellfish feed on suspended phytoplankton, which are trapped from water pumped across the gills by ciliary action. Many shellfish live in waters that receive runoff from agricultural, industrial, and residential areas and therefore Cryptosporidium oocysts may be filtered by the gills during feeding, and become concentrated in the digestive glands/tract. If these oocysts are not excreted or inactivated by the shellfish, or in subsequent preparatory processes, they may be ingested by consumers, the shellfish thereby acting as vehicles of infection.69 Oocysts can remain viable for up to one year in seawater and can be filtered out by benthic mussels in which they retain their infectivity.70 The prolonged survival of oocysts in seawater has important implications to coastal recreational waters and shellfish-rearing waters, contaminated with human and animal sewage. Cryptosporidium has been described in all the major food animals, including sheep, cattle, pigs, and poultry.11 During the slaughter of cattle and sheep, contamination of the carcass and edible offals with viable oocysts remains a major cause for concern. Unlike poultry processing, the dressing of the carcass in cattle and sheep slaughtering remains largely a nonautomated labor intensive operation. Contamination may occur through fecal spillage onto the hide or fleece, which may then contaminate the carcass

during removal.23 Whilst little information is available on contamination of fresh meat with oocysts, the high survival rate of the parasite in the environment, renders it likely that oocysts could persist in raw meat systems. A recent study examined the thermal inactivation of oocysts when attached to a beef surface.71 The authors have shown that C. parvum oocysts remain viable during mild heat treatments, i.e., low temperature/short time, but lose viability (as assessed by vital staining or HCT-8 infectivity rates) if subjected to higher temperatures and/or longer treatment times. Thus it appears that thermal processing, already widely applied to reduce the numbers of viable undesirable microorganisms (principally prokaryotes) in food products could be used to reduce viable C. parvum oocyst numbers in these complex matrices. There have been numerous studies reporting the ability of flies to act as mechanical vectors in the transmission of oocysts.72–82 Transmission of Cryptosporidium by adult flies occurs via (i) mechanical dislodgment from the exoskeleton; (ii) fecal deposition; and (iii) regurgitation.75 It has been demonstrated that filth flies can transport infectious oocysts of C. parvum on their external surfaces and in their digestive tracts.72–74 Exposure of adult house flies to bovine diarrheal feces contaminated with C. parvum resulted in intense deposition of the oocysts through fecal spots and vomit drops on the surfaces visited by house flies.73 A recent study in northwest Georgia (USA) detected Cryptosporidium in 55.56% of three different families of synanthropic flies.81 Cockroaches, dung beetles and birds have also been shown to carry the parasite and can be a source of contamination.83–85

Cryptosporidium

46.1.4 Methods for Diagnosis of Foodborne Cryptosporidium Detection of Cryptosporidium in food involves isolating the parasite from the foodstuff and then detection using a variety of methods including microscopy, immunofluorescent antibody staining, and other immunological detection methods including enzyme linked immunosorbent assay (ELISA’s) and DNA-based detection methods. Currently, there are no national or international guidelines for determining oocyst contamination in or on foodstuffs and isolation of the parasite from different food-types is problematic, as very different recovery efficiencies have been reported (see Section 46.1.4.3). 46.1.4.1 Conventional Methods for Detection of Cryptosporidium Traditionally, the detection of Cryptosporidium oocysts in environmental, water, food and fecal samples has primarily relied on examination by microscopy. However, morphological characters for identifying Cryptosporidium are few, and identification based on light microscopy alone is unreliable and relatively time consuming.86 When using conventional microscopy, differential staining techniques are usually required due to the fact that oocysts are similar in size and shape to yeasts, fecal components and other debris.87 A safranin-methylene blue stain has been described,88 which stains oocysts red and yeasts and other fecal debris blue. Acid-fast stains such as Kinyoun,89 ZiehlNeelsen,90 and DMSO-carbol fuchsin,91 stain oocysts red with the background counterstained and are commonly used in many laboratories. Differential staining techniques, however, are time consuming and vary in their sensitivity and specificity.88,92,93 In addition, unless samples have been fixed overnight in formalin, oocysts may frequently take up the counter stain. Negative staining techniques using nigrosin,94 light green and merbromide,95 malachite green,96 which stain background yeasts and bacteria but not the oocysts, have also been developed. Negative staining methods are faster but are considered by some to be less sensitive than conventional staining techniques.97,98 In general detection methods that rely on conventional microscopy lack sensitivity, are time consuming and require an experienced microscopist due to the resemblance of oocysts to background fecal debris, which can make identification difficult. Immunological detection methods offer superior speci­ ficity,99 and immunofluorescent antibody (IFA) staining has been widely used for the detection of Cryptosporidium in fecal smears, water and food.23,51,100,101 Both polyclonal and monoclonal antibodies (MAbs) have been raised against surface exposed epitopes on purified oocysts and fluorescently labeled antibody preparations are readily available commercially. A limitation of polyclonal immunodetection methods is that they suffer from cross reaction with other organisms.102

655

MAbs offer increased specificity and both direct and indirect detection methods are available. Commercially available FITC-MAbs are used routinely for detecting and enumerating oocysts in environmental samples.101,103 There are no commercially available antibody preparations specific for epitopes on human-pathogen Cryptosporidium oocysts. Different commercial MAbs recognize different sets of surface epitopes,104 and most, if not all commercially available MAbs have been raised against epitopes on a limited number of C. parvum isolates.101 Therefore if the FITC-MAbs supplied in the kit does not bind to a particular isolate due to antigenic variability of the oocyst wall epitopes, false negatives can occur. Oocysts of different species and genotypes of Cryptosporidium can also express a range of oocyst wall epitopes and differences in staining intensity has been observed in purified C. hominis and C. parvum oocyst isolates using three commercially available FITC-MAb preparations.101 False negatives can also occur due to the fact that the target epitopes can be easily stripped from the organism by mild environmental perturbations.23 In general, IgG MAbs appear to have increased avidity to oocyst surface antigens compared with IgM, allowing more efficient oocyst concentration, resulting in increased recoveries using immuno-magnetic separation (IMS) from water samples with high turbidity.105 IgG1 antibodies have been shown to achieve a better diagnostic specificity compared with IgG3 or IgM for the detection of Cryptosporidium oocysts in environmental samples.106,107 Enzyme linked immunosorbent assays (ELISA’s) and enzyme immunoassays (EIA’s) for Cryptosporidium are also available and have the advantage of reducing assay times and being amenable to automation. An immunochromatographic (dipstick) assay is also available. However, the sensitivity of all immunological-based methods is relatively low. For example, between 1×104 and 5×104 oocysts per gram of unconcentrated feces are necessary to obtain 100% detection efficacy using either conventional differential staining or a commercially available FITC-MAbs methods.108 The sensitivity of ELISA’s has been reported to be as low as 3×105–106 oocysts per milliliter.109 Biosensor chips that detect and quantitate C. parvum in real-time via anti-C. parvum IgM binding have been developed but have not been tested on food.110,111 A major limitation of all these techniques however, is their inability to speciate due to morphological similarity between many species. For example, oocysts of C. parvum, C. hominis, C. meleagridis and C. bovis have no apparent internal features that are unique among them and they overlap in size, ranging from 4.2–4.8 µm to 4.8–6.0 µm.1,112 46.1.4.2 Molecular Methods for Detection of Cryptosporidium Since the development of the polymerase chain reaction (PCR), PCR-based techniques have permitted specific and sensitive detection of Cryptosporidium spp. for clinical diagnosis and environmental monitoring.13,101 The advantages of PCR-based detection of Cryptosporidium in clinical

656

Molecular Detection of Foodborne Pathogens

and environmental samples are: its high sensitivity, specificity and rapidity enabling large numbers of samples to be analysed at a time; ease of use; relatively low cost and; ability to genotype, therefore, eliminating false-positives due to cross contamination and allowing the detection of the range of pathogens present.113,114 The sensitivity and specificity of PCR-based detection technique allow for the detection of a single oocyst in clinical and environmental samples.115–117 There is no standard locus recommended for detecting Cryptosporidium, and a wide variety of loci have been utilized (Table 46.4), but PCR-RFLP or sequence analysis of the 18S rRNA gene is the most appropriate method as, due to its conserved nature, it results in amplification of all Cryptosporidium species and genotypes. A nested PCR-RFLP 18S assay has been developed which is widely used and can detect most species and genotypes of Cryptosporidium.116,118 The primary PCR product produced by the external primers is 1325 bp long and the internal product is 826 bp. The Xiao et al. primers are specific for Cryptosporidium but as they amplify a relatively large PCR product, samples that contain very low numbers of oocysts, or partially degraded DNA, will not always be amplified. As the numbers of oocysts extracted from foodstuffs is typically low, primers which amplify a smaller 18S rRNA PCR product offer more sensitivity.119 PCR tools based on either antigen genes, or housekeeping genes such as the heat shock protein 70 (HSP70), or the Cryptosporidium oocyst wall protein (COWP) gene, can sometimes fail to detect the DNA of some of the human-pathogenic species including C. felis, C. canis, and C. muris, and are therefore more effectively

used as secondary tools for detecting common species such as C. hominis and C. parvum.120 Real-time quantitative PCR (RT-qPCR), has overcome several limitations of the conventional PCR and allows for the quantitation of the number of starting copies of a specific template in the sample. In RT-qPCR the amplified product is detected using fluorescent dyes. These fluorescent dyes are linked to oligonucleotide probes and/or primers, which bind specifically to or are incorporated into, the amplified PCR product. Changes in the fluorescence intensities are monitored during the PCR reaction (in “real-time”) during the exponential phase. The threshold cycle (or Ct) when product is first detected is compared with a standard curve (based on Ct value versus known concentrations of DNA standards) to estimate the amount of material in the test sample. In addition, the whole process is faster than regular PCR as no electrophoresis is required reducing the possibility for cross contamination and false-positive results.138 Quantitative real-time PCR detection methods have been developed to quantitate the numbers of Cryptosporidium oocysts present in fecal and water samples.114,139–142 QPCR can also be coupled with high resolution melt (HRM) curve analysis to differentiate different species of Cryptosporidium. HRM curve analysis involves the gradual dissociation of relatively AT-rich regions (“lower” melting domains) within the amplicon prior to the ultimate melting. Single nucleotide polymorphisms or heteroduplexes can result in slight but measurable changes in the HRM profile by affecting amplicon dissociation.142,143 The recent development of subgenotyping tools is important for foodborne detection as it facilitates tracking of the

Table 46.4 Some of the Loci used to Detect Cryptosporidium Locus Acetyl-CoA Actin Beta-tubulin COWP COWP DHFR DHFR DHFR HSP-70 ITS1 of rRNA Poly-T TRAP-C1 18S rRNA 18S rRNA 18S rRNA 18S rRNA 18S rRNA 18S rRNA Unknown Unknown

Product Size (bp)

Type of PCR

Reference

357 (C. hominis), 190 (C. parvum) 1950 513 (C. hominis), 519 (C. parvum) 318 1200 556 445 1750 ~825 590–593 597 1353 C. hominis (411), 311 (C. parvum)

Nested PCR Nested PCR PCR PCR-RFLP Nested PCR Nested PCR-RFLP Nested PCR Species-specific PCR Nested PCR Species-specific PCR PCR-RFLP PCR-RFLP PCR-RFLP PCR PCR-RFLP Nested PCR-RFLP Nested PCR-RFLP Nested PCR PCR-RFLP Species-specific PCR

121 122 123 124 125 126 127 128 129 130 131 132 133 134 135 116 117 119 136 137

347 1095 ~592 553 1033 408

Cryptosporidium

source of the infection.144,145 The term “subgenotype” refers to genetic differences within Cryptosporidium species/ genotypes (i.e., variation within C. hominis or C. parvum etc.).18,145 Subgenotype analysis is more informative than genotyping because it can elucidate genetic variation on a finer scale and is therefore more accurate to track contamination sources in outbreak investigations or contamination of food. Subgenotyping is also important in assessing the zoonotic potential of the parasite, as it has been shown that some subtypes of C. parvum only circulate within the human population.18,145 Currently, there are several subgenotyping tools developed for the genetic characterization of Cryptosporidium including microsatellite analysis, and sequence analysis of the GP60 gene, the HSP70 gene, a double-stranded (ds) RNA virus that is present in most Cryptosporidium species, a 47-kDa protein (CP47), a mucin-like protein (Mucin1), a serine repeat antigen (MSC6-7), and a 56-kDa trans-membrane protein (CP56) in chromosome 6, the 70 kDa heat shock protein (HSP70) in chromosome 2, and a T-rich gene fragment (Chrom3T) in chromosome 3.18,146 Microsatellites are short sequence repeats that constitute a rich source of polymorphisms, which has subsequently led them to be used extensively for subgenotyping/ fingerprinting.147–150 Microsatellites located in noncoding regions are more likely to accumulate mutations than protein coding sequences, and have been therefore chosen for subgenotyping studies.148 A recent international interlaboratory comparative trial revealed that GP60 subgenotyping is a more reliable method than microsatellite analysis as a fingerprinting tool for Cryptosporidium.144 The GP60 gene, also known as Cpgp15/45, exhibits a high degree of sequence polymorphism, far greater than any other coding Cryptosporidium locus examined to date.151,152 It is a single copy gene encoding a precursor protein that is proteolytically cleaved during the intracellular stages of the parasite to yield mature cell surface glycoproteins, known as gp45 and gp15. Both glycoprotein products are expressed on the apical and surface regions of the invasive stages of the Cryptosporidium lifecycle and have been implicated in the attachment to and invasion of host cells.151–153 There are a number of significant and epidemiologically important advantages of using sequence analysis of the GP60 gene including (i) the high degree of sequence polymorphism; (ii) the full comparability of data across laboratories; (iii) the availability of data from several waterborne and foodborne outbreaks; and (iv) the biological relevance.151,152 Sequence analysis of the GP60 gene has identified a number of subgenotypes, associated with the C. parvum and C. hominis species (11 and nine respectively), that show variations throughout the entire length of the gene, and which have been found to display the most sequence diversity within a TCA/TCG trinucleotide repeat region that is present in all GP60 sequences.152,154,155 Cryptosporidium hominis subgenotypes are designated Ia, Ib etc., whereas C. parvum subgenotypes are designated IIa, IIb etc. There are also further “subtypes” within each subgenotype, which

657

mostly differ from each other in the number of TCA repeats, and each subgenotype may contain more than one subtype.13,154,155 C. parvum IIa subtypes are commonly found in human and animals. However, GP60 analysis has revealed that the detection of C. parvum in humans does not always indicate a zoonotic source of infection, as a humanadapted C. parvum subgenotype (IIc) has not been reported in animals.13,145 46.1.4.3 Detection of Cryptosporidium in Fruit and Vegetables Standardized methods for detecting Cryptosporidium on soft fruit and salad vegetables are not available. In 2004, Ripabelli and colleagues investigated the recovery of C. parvum oocysts from experimentally contaminated lettuce leaves.3 Material recovered from the lettuce samples by washing in detergent solutions were concentrated by filtration using the Envirochek sampling capsule. Oocysts were concentrated by immunomagnetic separation (IMS) and detected by microscopy following modified Ziehl–Neelsen (MZN) staining. Parasite DNA was detected using microscopy and a nested-PCR assay of the COWP gene. However, after IMS, the recovery rates were very low (0–6.5%). More recently Cook and colleagues optimized and validated methods for the recovery of Cryptosporidium on lettuce and raspberries.156,157 The method involves extraction of oocysts from the foodstuffs, concentration and separation of the oocysts from food materials and fluorescent staining of the oocysts followed by fluorescent microscopy. Extraction of oocysts from the foodstuffs involved addition of a liquid extractant to the sample, followed by orbital shaking or rolling to facilitate removal of the oocysts from the food surface. The extract is then concentrated by centrifugation, before IMS of the oocysts from the residual food materials. Recoveries of 59.0±12.0% of C. parvum oocysts seeded onto lettuce, and of 41.0±13.0% of C. parvum oocysts seeded onto raspberries (n=30 for each matrix) were reported.156 This was followed up with an interlaboratory collaborative trial involving eight expert laboratories in the United Kingdom.157 Results revealed that the method was just as reproducible between laboratories, as it was within a single laboratory. The total median percentage recovery (from all artificially contaminated samples) produced by the method was 30.4%.157 Since the experimentally determined lowest ID50 for Cryptosporidium in human volunteers is nine oocysts,158 the above methodology has the required sensitivity for detecting Cryptosporidium in foodstuffs. Few studies have utilized PCR detection of Cryptosporidium on fruit and lettuce. Ripabelli and colleagues compared microscopy and a nested-PCR assay of the COWP gene. Detection by PCR was less sensitive than microscopy.3 A nested PCR amplification of the 18S rRNA gene identified one Cryptosporidium positive in a variety of lettuce tested (n=50; iceberg and round) at retail sale in Northern Ireland.30 Sequence analysis of 361 bp showed 100% homology with C. parvum and/or C. hominis indicating the potential for human infection.30

658

46.1.4.4 Detection of Cryptosporidium in Meat, Fish, and Shellfish As with vegetables, standardized methods for detecting Cryptosporidium in/on meat, fish, and shellfish are not available. Little is known about the prevalence of Cryptosporidium on raw meat and fish. Recently, acute gastroenteritis was reported among four members from the same company who had eaten a raw meat dish called “Yukke: Korean-style beef tartar” and raw liver at a rotisserie in Sakai City, Japan.159 Stool specimens obtained from three of the four symptomatic patients were positive for C. parvum after PCR analysis at the 18S locus, and subtyping at the GP60 loci identified the IIa subtype.159 The prevalence of Cryptosporidium in shellfish varies widely and prevalences of 6–55.6% have been reported.69 True prevalences are difficult to determine as recovery efficiencies from shellfish tend to be variable, ranging from 12% to 70%.36,160,161 It is also believed that the consistency of the shellfish tissue affects the performance of the IMS, being less effective in the more mucoid shellfish. There is also debate as to which tissue should be used for analysis, however most researchers use tissue homogenates rather than gill samples (homogenate or washings) or hemolymph when analyzing for Cryptosporidium.69,162 Molecular characterization of Cryptosporidium oocysts detected in marine bivalves, have reported a preponderance of C. parvum, although other species have also detected (including C. meleagridis, C. bailieyi, C. hominis, C. felis and C. andersoni).69 46.1.4.5 Detection of Cryptosporidium in Drinks Outbreaks of illness associated with consumption of fruit juice have been a growing public health problem since the early 1990s.163 In the US, from 1995 to 2005, 21 juiceassociated outbreaks were reported to Centers for Disease Control (CDC); ten implicated apple juice or cider, eight were linked to orange juice, and three involved other types of fruit juice. These outbreaks caused 1,366 illnesses, with a median of 21 cases per outbreak (range, two to 398 cases).163 Methods for detecting Cryptosporidium in drinks are similar to those for the recovery and detection of parasites in water and are usually based on the USEPA 1623 method (US Environmental Protection Agency, 2001), which uses filtration, followed by elution, concentration by IMS, and detection by IFA. Sheather’s sucrose gradient centrifugation has also been used to concentrate oocysts from apple cider.2 Molecular analysis based on the COWP locus has been used to detect one to ten oocysts in 20 ml of artificially contaminated raw milk.164 Cryptosporidium DNA from 17 different isolates was used to spike milk samples and screened by PCR analysis of a 358-bp fragment of a COWP gene.164 By combining PCR amplification and DNA hybridization, the sensitivity of the assay was increased to the femtogram range but the primer sets/probes utilized were unable to detect all isolates of Cryptosporidium used.164 PCR methods have also been employed to detect Cryptosporidium in/on raw unwashed apples, washed apples, cleaning water, fresh cider, and finished cider samples were collected from five Ontario

Molecular Detection of Foodborne Pathogens

producers of cider.165 Cryptosporidium was detected in 2/111 water samples, 1/114 washed apple samples, 5/113 fresh cider samples, and 2/113 finished cider.165 Molecular and epidemiologic methods were used to link ozonated apple cider to an outbreak of cryptosporidiosis in 2003 in Ohio.50 PCR analysis of 14 available samples from the laboratory-confirmed case-patients identified 12 positives; 11 of these were C. parvum and remaining case was the cervine Cryptosporidium genotype (W4). Subtype identification of stool samples at the GP60 locus identified the C. parvum subtypes (IIaA15G2R1 and IIaA17G2R1). The remaining contents of a jug of cider that a laboratory-confirmed casepatient had partially drunk were also positive by PCR for C. parvum subtype IIaA17G2R1. This case-patient’s stool sample yielded the same subtype of C. parvum, as did four other case-patients’ stool samples; all of these persons drank cider in the two weeks before illness onset.50 The two C. parvum subtypes found in this outbreak likely represent a common contamination source; both are common in cattle, and multiple subtypes are commonly found on farms.13 The study also revealed that ozonation against Cryptosporidium in food or juice products was inadequate to kill Cryptosporidium. The study resulted in the FDA issuing an addendum to its HACCP rule that advised that juice makers should not use ozone in their manufacturing process unless they can prove a 5-log pathogen reduction through ozonation.50 A recent study compared two techniques for preparing DNA templates, following IMS purification: a filter-based method using the FTA Concentrator-PS filter (Whatman, Inc., Newton, MA) and a total DNA extraction method using a MasterPure DNA kit by Epicentre (Madison, WI).4 Six foods (orange juice, apple cider, whole milk, strawberries, parsley, and lettuce) were artificially contaminated with known numbers of oocysts. Following IMS purification, DNA templates were prepared from acid-eluted oocysts using the two methods above and Cryptosporidium DNA was detected using a nested PCR assay targeting the 18S rDNA gene.117 Results revealed that FTA Concentrator-PS units were less sensitive and introduced replicate variability with lower levels of inocula (50 and five oocyst samples) than the conventional DNA extraction kit. However the FTA Concentrator-PS units required considerably less time and fewer sample manipulations.4 Of the different food types tested, cider was considered the most difficult. Other groups have also reported PCR inhibition, most likely due to polyphenolics, when attempting to detect pathogens in apple cider.2,166 Another study has compared the effectiveness of six DNA extraction methods for the detection of Cryptosporidium with oocyst-seeded samples, DNA-spiked samples, and field water samples.167 The study also evaluated the effects of different PCR facilitators for the relief of inhibitors of PCR amplification. The results of seeding and spiking studies showed that PCR inhibitors were presented in all DNA solutions extracted by the six methods. However, with the inclusion of bovine serum albumin in the PCR mixture, DNA extracted with the FastDNA SPIN kit for soil without oocyst isolation resulted in PCR performance similar to that produced by the QIAamp

659

Cryptosporidium

DNA minikit after oocysts were purified by immunomagnetic separation.

46.2  Methods 46.2.1 Reagents and Equipment The reagents and equipment that are required for extraction of Cryptosporidium DNA and subsequent PCR amplification are listed on Table 46.5.

46.2.2 Sample Preparation For the recovery of Cryptosporidium oocysts from vegetables and lettuce, the following methodology described by Cook et al.,156 appears to provide the most reproducible results (see Table 46.5).157 (1) Place food sample (30 g) in a clean 250 ml glass beaker and add then 150 ml of 0.1 M glycine. (2) Place beaker on an orbital shaker and shake at low speed (80 rpm) for 1 min. (3) Strain contents of filter through a filtered Stomacher® bag (Seward, UK) into another clean 250 ml beaker. (4) Rinse out the beaker with 50 ml of 0.1 M glycine. (5) Divide extract between four 50 ml centrifuge tubes, and centrifuge at 2,500×g for 10 min at 15°C (4°C for raspberry samples). (6) Decant supernatants into a clean glass beaker, leaving approximately 1 ml fluid with each pellet. Table 46.5 Reagents, Supplies and Equipment Needed for Extraction of Cryptosporidium DNA and Detection via PCR Amplification Reagents 1 M glycine pH 5.5 dH2o Anti-Cryptosporidium magnetic bead kit PCR grade H2o Nonacetylated BSA Primers (see Section 46.2.3) Taq polymerase+associated buffers dNTP’s Restrictions enzymes (Vsp I, Ssp I, Dde I) Loading buffer and molecular weight markers

Supplies and Equipment Filtered Stomacher® bag Orbital shaker 250 ml beakers Leighton tubes Disposable gloves Pipette tips: 10 μl, 100 μl, 1000 μl Tubes: 500 ml, 50 ml, 10 ml, 2 ml, 1.5 ml, 0.1 ml Refrigerated bench-top centrifuge Rotating wheel Magnet DNA extraction kits (Qiagen stool kit, Qiagen Dneasy Tissue Kit, FastDNA SPIN kit). Thermocycler Horizontal electrophoresis apparatus

(7) Combine pellets by pastette into a Leighton tube (ImmuCell Corporation, Portland, ME). (8) Rinse centrifuge tube with approximately 3 ml of the decanted supernatant, serially transferred from tube to tube, and then add to the Leighton tube. (9) Repeat washing procedure until the volume in the Leighton tube is approximately 10 ml. (10) Resuspend pellet in 0.5 ml by vortexing, prior to IMS. (11) Separate oocysts from pellet using IMS according to the manufacturer’s protocol depending on the brand of IMS used. (12) If using Dynabeads (Dynal, Lake Success, NY), resuspend pellet in 0.5 ml and then add 1 ml of 10× buffer A, 1 ml of 10× buffer B, and 100 µl of antiCryptosporidium Dynabeads from the Dynabead anti-Cryptosporidium kit (Dynal) to a 15-ml tube containing 0.5 ml of pellets and adjust the final volume to 10 ml. (13) Rotate tube at 20 rpm at room temperature for 1 h. (14) Capture Cryptosporidium oocysts using a magnetic device (MPC; Dynal) and use the Dynabead/oocyst mixture directly in DNA extraction without oocyst detachment. For extraction of oocysts from water and drinks, the standard USEPA 1623 method should be used (www.epa.gov/ waterscience/methods/1623.pdf). This involves filtering 10 L (or less) of water/drink through a Pall Gelman Envirochek™ HV filter (or a variety of other EPA approved filtration methods) according to the manufacturers instructions. For DNA extraction, various methods have been compared and the QIAamp DNA stool minikit (Qiagen) and the FastDNA SPIN kit (BIO 101) give comparable results.167 The advantage of using the FastDNA SPIN kit is that DNA can be extracted from water/food samples following filtration, without the IMS step, which greatly reduces analysis time. The addition of 400 ng/µl of nonacetylated BSA (Sigma, St. Louis, MO) to the PCR results in significant reduction of PCR inhibitors. The use of the FastDNA SPIN kit, without the IMS step has not been tested on food samples. For DNA extraction using the QIAamp DNA stool minikit, extract DNA from Dynabead/oocyst mixture by adding 180 µl of the ATL buffer from the QIAamp DNA minikit, transfer the suspension into a 1.5-ml Eppendorf tube and subject to five cycles of freeze-thaw (–70°C for 30 min and 56°C for 30 min). Extract DNA from the purified oocysts with the QIAamp DNA minikit using the manufacturerrecommended procedures. If using the FastDNA SPIN kit for extraction from water/ drinks, re-suspend filtered oocysts in 0.5 ml and transfer into a 2-ml tube containing the lysing matrix E from the FastDNA SPIN kit. Add 978 µl of sodium phosphate buffer and 122 µl of MT buffer from the kit and vortex in a FastPrep instrument (BIO 101) for 30 sec at the speed setting 5.5. Process samples further in accordance with the manufacturer-recommended procedures.

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Molecular Detection of Foodborne Pathogens

46.2.3  Detection Procedures 46.2.3.1 Nested PCR Targeting a 825-bp 18S rDNA Fragment Principle: A two-step nested PCR protocol was described by Xiao et al.,116 to amplify an 826 bp region the 18S rDNA gene. Procedure: (1) Primary PCR: a PCR product of about 1,325 bp is amplified using primers 5′-TTCTAG AGCTAA TACATGCG-3′ and 5′-CCCATTTCCTT CGAAA CAGGA-3′. PCR reaction components: water, 10 μl of 10× PCR buffer, 6 mM MgCl2, 200 μM (each) deoxynucleoside triphosphate (dNTP), 100 nM (each) primer, 2.5 U of Taq polymerase, and 0.25–1 μl of DNA template in a total 100-μl reaction mixture. Amplification conditions: A total of 35 cycles, each consisting of 94°C for 45 s, 55°C for 45 s, and 72°C for 1 min, with an initial hot start at 94°C for 3 min and a final extension at 72°C for 7 min. (2) Secondary PCR: a secondary PCR product of 825– 864 bp (depending on the species/genotype) is then amplified from 2 μl of the primary PCR with primers 5′-GGAAGGGTTGTATTTATTAGATAAAG3′ and 5′-AAGGAGTAAGGAACAACCTCCA-3′. The PCR and cycling conditions are identical to primary PCR, except that 3 mM MgCl2 is used. (3) RFLP analysis: 20 μl of the secondary PCR product is digested in a total of 50 μl of reaction mixture,

consisting of 20 U of SspI or VspI and 5 μl of respective restriction buffer at 37°C for 1 h, under conditions recommended by the supplier. The digested products should then be fractionated on 2.0% agarose gels. Note: Table 46.6 lists the predicted RFLP fragment sizies for some species and genotypes of Cryptosporidium. VspI digestion is required to differentiate between C. parvum and C. hominis parasites. Because C. andersoni and C. muris had identical SspI and VspI restriction patterns, they can be differentiated by digesting secondary PCR products with 20 U of DdeI at 37°C for 1 h under conditions recommended by the supplier.118 46.2.3.2 Nested PCR Targeting a 587-bp 18S rDNA Fragment Principle: A two-step nested PCR protocol was described by Ryan et al.,119 to amplify a 587 bp fragment of the 18S rDNA gene. Procedure: (1) Primary PCR: a PCR product of 763 bp is amplified using the forward primer 18SiCF2 (5′-GAC ATA TCA TTC AAG TTT CTG ACC-3′) (bp position 292) and the reverse primer 18SiCR2 (5′-CTG AAG GAG TAA GGA ACA ACC-3′) (bp position 1007). PCR reaction components: water, 200 µM each of dNTP, 1× PCR buffer, 1.5 mM MgCl2, 0.5 units of Taq polymerase and 12.5 pmoles of forward and reverse primers in a total of 25 µl reaction. PCR

Table 46.6 Predicted RFLP Fragment Sizes Following Digestion with Ssp I and/or Vsp I for Various Cryptosporidium Species and Genotypes Species

SspI Digestion (bp)

VspI Digestion (bp)

Source

PCR Fragment (bp)

C. andersoni C. serpentis C. baileyi C. felis C. meleagridis C. wrairi C. varanii C. hominis C. parvum

Cattle, camel, hyrax Snake Chicken Cat Turkey Guinea pig Desert monitor Human Bovine

833 831 826 864 833 834 834 837 834

385, 448 14, 33, 370, 414 254, 572 15, 33, 390, 426 11, 11, 108, 254, 449 11, 11, 109, 254, 449 19, 33, 109, 255, 418 11, 12, 111, 254, 449 11, 12, 108, 254, 449

102, 731 102, 729 102, 104, 620 102, 104, 182, 476 102, 104, 171, 456 102, 104, 628 102, 104, 628 70, 102, 104, 561 102, 104, 628

Type A C. parvum

Bovine

831

9, 119, 254, 449

102, 104, 625

Type B Mouse genotype I C. canis Ferret genotype C. suis C. fayeri

Mouse Dog Ferret Pig Kangaroo, koala

838 829 837 838 837

11, 12, 112, 254, 449 20, 33, 105, 254, 417 11, 12, 111, 254, 449 9, 11, 365, 453 33, 109, 254, 441

102, 104, 175, 457 94, 102, 633 102, 104, 174, 457 102, 104, 632 102, 104, 631

Source: Modified from Xiao, L. et al., Appl. Environ. Microbiol., 65, 1578, 1999.

661

Cryptosporidium

amplification: Forty-five PCR cycles (94°C for 30 sec, 58°C for 30 sec, 72°C for 30 sec) with an initial hot start (94°C for 5 min) and a final extension (72°C for 10 min). (2) Secondary PCR: a fragment of ~587 bp is amplified using 1 µl of primary PCR product and the nested forward 18SiCF1 (5′-CCT ATC AGC TTT AGA CGG TAG G-3′) (bp position 289) and nested reverse 18SiCR1 (5′-TCT AAG AAT TTC ACC TCT GAC TG-3′) (bp position 851) primers. The PCR conditions for the secondary PCR are identical to the primary PCR. Sequence analysis is then used to identify to species/genotype level. 46.2.3.3 Nested PCR Targeting the GP60 Locus Principle: A two-step, nested PCR for the GP60 locus works well.151,168

potentially used in bioterrorist attacks and is categorized as NIH category B biodefense agent (http://www.niaid.nih.gov/ biodefense/bandc_priority.htm). With the development of improved laboratory detection systems for isolation, identification and genotyping, coupled with food-related outbreaks, more attention is being placed on the potential transmission of Cryptosporidium through foodstuffs.23 Thus, food-testing laboratories will experience an increased demand for having such assays in place to routinely monitor for this organism, to evaluate its viability, and, ideally, to distinguish between human and nonhuman pathogens in order to estimate the public health risk posed by food contamination. In view of the large spectrum of food matrices on which analytical methods should be applied, more work is needed to evaluate the performance of the available techniques, particularly to improve the recovery rates, and to develop rapid and cost-effective molecular methods for the identification of human pathogens on foodstuffs.

Procedure: (1) Primary PCR: a PCR product of ~995 bp is amplified using the forward primer, StrongGP60 F1 (5′-ATGAGATTGTCGCTCATTATC-3′), and the reverse primer, StrongGP60 R1 (5′-TTACAACACG AATAAGGCTGC-3′).151 PCR reaction components: 0.01–1.0 ng of DNA should be amplified in a total volume of 25 µl reaction mixture consisting of water, 0.5 U of Tth polymerase, 10× DNA reaction buffer, 1.5 mM MgCl2, 0.2 mM of each dNTP and 12.5 pmol of forward and reverse primers. Amplification conditions: a preliminary cycle of 94°C for 2 min, 56°C for 1 min and 72°C for 2 min, followed by 45 PCR cycles of 94°C for 30 sec, 56°C for 20 sec and 72°C for 1 min, and a final extension of 72°C for 7 min. (2) Secondary PCR: the primers CDCGP60 F2 (5′-TCC GCTGTATTCTCAAGCC-3′) and CDCGP60 R2 (5′-GCAGAGGAACCAGCATC-3′) are used to amplify a fragment of ~832 bp.168 The secondary PCR is set up as for the primary PCR. The secondary PCR cycling conditions are identical to that of the primary reaction, with the exception that the annealing temperature is reduced from 56°C to 54°C. Sequence analysis is then used to identify to subtype level.

46.3 Conclusion and Future Perspectives Cryptosporidium has emerged as an important gastrointestinal parasite of humans and animals, with a global distribution. The oocysts produced by Cryptosporidium can survive for several months in the environment, are easily spread via water, and are difficult to inactivate or remove from water intended for consumption without the use of filtration. For these reasons, cryptosporidiosis represents the major public health concern of water utilities in developed nations.11,13 It is also one of the few foodborne parasites that can be

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665 151. Strong, W.B., Gut, J. and Nelson, R.G. Cloning and sequence analysis of a highly polymorphic Cryptosporidium parvum gene encoding a 60-kilodalton glycoprotein and characterization of its 15- and 45-kilodalton zoite surface antigen products. Infect. Immunol., 68, 4117, 2000. 152. Alves, M. et al. Subgenotype analysis of Cryptosporidium isolates from humans, cattle, and zoo ruminants in Portugal. J. Clin. Microbiol., 41, 2744, 2003. 153. Leav, B.A. et al. Analysis of sequence diversity at the highly polymorphic Cpgp40/15 locus among Cryptosporidium isolates from human immunodeficiency virus-infected children in South Africa. Infect. Immunol., 70, 3881, 2002. 154. Peng, M.M. et al. A comparison of Cryptosporidium subgenotypes from several geographic regions. J. Euk. Microbiol., Suppl, 28S, 2001. 155. Sulaiman, I.M. et al. Unique endemicity of cryptosporidiosis in children in Kuwait. J. Clin. Microbiol., 43, 2805, 2005. 156. Cook, N. et al. Towards standard methods for the detection of Cryptosporidium parvum on lettuce and raspberries. Part 1: development and optimization of methods. Int. J. Food. Microbiol., 109, 215, 2006. 157. Cook, N. et al. Towards standard methods for the detection of Cryptosporidium parvum on lettuce and raspberries. Part 2: validation. Int. J. Food. Microbiol., 109, 222, 2006. 158. Okhuysen, P.C. et al. Virulence of three distinct Cryptosporidium parvum isolates for healthy adults. J. Infect. Dis., 180, 1275, 1999. 159. Yoshida, H. et al. An outbreak of cryptosporidiosis suspected to be related to contaminated food, October 2006, Sakai City, Japan. Jpn. J. Infect. Dis., 60, 405, 2007. 160. Gómez-Couso, H. et al. Levels of detection of Cryptosporidium oocysts in mussels (Mytilus galloprovincialis) by IFA and PCR methods. Vet. Parasitol., 141, 60, 2006. 161. MacRae, M. et al. The detection of Cryptosporidium parvum and Escherichia coli O157 in UK bivalve shellfish. J. Microbiol. Methods, 60, 395, 2005. 162. Downey, A.S., and Graczyk, T.K. Maximizing recovery and detection of Cryptosporidium parvum oocysts from spiked eastern oyster (Crassostrea virginica) tissue samples. Appl. Environ. Microbiol., 73, 6910, 2007. 163. Vojdani, J.D., Beuchat, L.R. and Tauxe, R.V. Juice-associated outbreaks of human illness in the United States, 1995 through 2005. J. Food Prot., 71, 356, 2008. 164. Laberge, I. et al. Detection of Cryptosporidium parvum in raw milk by PCR and oligonucleotide probe hybridization. Appl. Environ. Microbiol., 62, 3259, 1996. 165. Garcia, L. et al. Potential sources of microbial contamination in unpasteurized apple cider. J. Food Prot., 69, 137, 2006. 166. Ogunjimi, A.A., and Choudary, P.V. Adsorption of endogenous polyphenols relieves the inhibition by fruit juices and fresh produce of immuno-PCR detection of Escherichia coli O157:H7. FEMS Immunol. Med. Microbiol., 23, 213, 1999. 167. Jiang, J. et al. Development of procedures for direct extraction of Cryptosporidium DNA from water concentrates and for relief of PCR inhibitors. Appl. Environ. Microbiol., 71, 1135, 2005. 168. Peng, M.M. et al. Molecular epidemiology of cryptosporidiosis in children in Malawi. J. Euk. Microbiol., 50 (Suppl), 557, 2003.

47 Cyclospora

Dongyou Liu, G. Todd Pharr, and Frank W. Austin Mississippi State University

Contents 47.1 Introduction ................................................................................................................................................................... 667 47.1.1 Classification and Biology ............................................................................................................................... 667 47.1.2 Clinical Presentation and Epidemiology ......................................................................................................... 668 47.1.3 Diagnosis ......................................................................................................................................................... 669 47.1.3.1 Phenotypic Techniques.................................................................................................................... 669 47.1.3.2 Molecular Techniques...................................................................................................................... 669 47.2 Methods.......................................................................................................................................................................... 671 47.2.1 Sample Preparation .......................................................................................................................................... 671 47.2.1.1 Sample Processing .......................................................................................................................... 671 47.2.1.2 DNA Isolation ................................................................................................................................. 671 47.2.2 Detection Procedures........................................................................................................................................ 671 47.2.2.1 PCR-RFLP . .................................................................................................................................... 671 47.2.2.2 Multiplex PCR ................................................................................................................................ 672 47.2.2.3 Real-time PCR ................................................................................................................................ 673 47.3 Conclusions and Future Perspectives............................................................................................................................. 673 References.................................................................................................................................................................................. 674

47.1  Introduction 47.1.1 Classification and Biology The genus Cyclospora covers a group of protozoan species in the kingdom Protista, phylum Apicomplexa, subclass Coccidiasina, order Eucoccidiorida and family Eimeriidae. As coccidian organisms, the members of Cyclospora genus resemble those of Cryptosporidium, Isospora, and Sarcocystis genera morphologically and biologically (see Chapters 46, 51, and 52). However, Cyclospora oocysts harbor two sporocysts each containing two sporozoites, in comparison to Cryptosporidium (whose oocysts have no sporocyst but four sporozoites), Isospora and Sarcocystis (whose oocysts all have two sporocysts each containing four sporozoites). In addition, Cyclospora oocysts (measuring between 8–36×7–22 μm in diameter, and C. cayetanensis oocyst measuring 8×10 μm) are larger than those of Cryptosporidium (5.2×4.9 μm), but smaller than those of Isospora belli (30×12 μm in diameter) and Sarcocystis (30×15 μm).1 Although the first Cyclospora organism was described in 1870 from intestines of the mole Talpa europaea, the genus Cyclospora was not established until 1881 to refer to C. glomericola observed in the millipede.2 For a long period, Cyclospora species have been considered as parasites of invertebrates, and attracted little attention. However, following the reports of undescribed coccidian diarrhea in Papua New Guinea in the 1970s, traveler’s diarrhea in Haiti, Nepal, and Peru in the 1980s, and chronic diarrhea in

acquired immunodeficiency syndrome patients in the 1990s, the importance of Cyclospora as a human pathogen has been recognized.3 With the early epidemiological and taxonomic work at the Universidad Peruana Cayetano Heredia in Lima, Peru, the Cyclospora species that is infective to humans is named Cyclospora cayetanensis.4 Among the 18 Cyclospora species that are identified to date, most are found in insetcts, reptiles, and mammals, C. cayetanensis is the only one capable of causing cyclosporiasis in humans (Table 47.1).1,5,6 Morphologically, the C. cayetanensis oocysts are 8–10 µm in size and spherical with a characteristic blue or green outer ring-fluorescence when examined under ultraviolet (UV) light, wherea oocysts from other species within the Cyclospora genera are larger and in different shapes. Like other coccidian pathogens, C. cayetanensis is an obligate intracellular parasite that completes its lifecycle within the host’s epithelial cells and gastrointestinal (GI) tract where it undergoes both asexual (merogony or schizogony) and sexual (gametogony) reproduction. Sporulated oocysts of C. cayetanensis are ingested by the human host through contaminated food or water. In the small bowl (typically the jejunum) the oocysts excyst with the assistance of bile, trypsin and other proteolytical factors in the tract. Crescent shaped sporozoites (of 1.2×9.0 μm in size) are then released. After gaining entry to epithelial cells of the small intestine, sporozoites undergo asexual reproduction (merogony) in parasitophorous vacuoles within epithelial cells to generate large numbers of merozoites. Apart from having the capacity to 667

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Molecular Detection of Foodborne Pathogens

47.1.2 Clinical Presentation and Epidemiology

Table 47.1 Cyclospora Species Identified to Date Species C. glomericola C. caryolytica C. viperae C. babaulti C. scinci C. tropidonoti C. zamenis C. niniae C. talpae C. megacephali C. ashtabulensis C. parascalopi C. angimurinensis C. cayetanensis C. colobi C. cercopitheci C. papionis C. schneideri

Main Host

Year of First Description

Insect Insect Reptile Reptile Reptile Reptile Reptile Reptile Insect Mammal/insect Mammal/insect Mammal Mammal/rodent Primate (Human) Primate Primate Primate Reptile

1881 1902 1923 1924 1924 1924 1924 1965 1968 1988 1989 1989 1990 1994 1999 1999 1999 2005

Source: Adapted from Shields, J.M. and Olson, B.H., Int. J. Parasitol., 33, 371, 2003 and Lainson, R., Mem. Inst. Oswaldo Cruz, 100, 103, 2005.

infect a new host cell and initiate another cycle of merogony, the merozoites can also proceed to sexual production in parasitophorous vacuoles via gametogony, by which merozoites turn into both male macrogamonts (which subsequently produce many microgametes) and female macrogamonts. The fusion between male microgamete and female macrogamont results in the formation of zygote that then matures into an oocyst. After disruption of the host cell, the oocyst is passed with the stool. Unlike the oocyst of Cryptosporidium, which exits the host as fully sporulated oocysts and is immediately infective, the oocyst of Cyclospora passed in the stool is unsporulated, and not infectious. The Cyclospora oocysts can survive in water at 4°C for 2 months and at 37°C for 7 days.7,8 To become infectious, Cyclospora oocyst needs to go through sporulation process (with its content i.e., sporont, dividing into two sporocysts, each containing two bananashaped sporozoites), which takes 7–13 days at temperatures between 22 and 32°C. However, sporulation of Cyclospora oocyst can be induced with 2.5% potassium dichromate, which is commonly used to induce sporulation in other coccidians. Alternatively, physical rupture of Cyclospora oocyst wall and exposure to 0.5% trypsin and 1.5% sodium taurocholate in PBS may also speed up the sporulation process. Even though Isospora belli oocyst is also shed as unsporulated form in human stool, it differs from Cyclospora oocyst in size and shape as well as in the number of sporocysts (two) and sporozoites (four) after sporulation.

Clinically, C. cayetanensis infections in humans often display GI symptoms such as loose or watery diarrhea, nausea, vomiting, and abdominal cramps accompanied by loss of appetite, and unintentional weight loss.9,10 In addition; human cyclosporiasis may present some nonspecific clinical signs such as fever, chills, muscle aches, joint aches, generalized body aches, headache, or fatigue.11–13 Although a case of C. cayetanensis infection is usually defined as diarrhea with three or more loose stools per day, asymptomatic infections or infection without watery diarrhea are known to occur. The onset of symptoms in endemic countries usually begins approximately 5–8 days (with a median incubation period of 7.5 days and range of 1–23 days) after infection and the symprtoms may persist for 1 month to 6 weeks. The severity and length of the illness varies significantly in relation to the age and immunological condition of the host.14 The potential complications associated with cyclosporiasis include Guillain–Barré syndrome, reactive arthritis syndrome (formerly Reiter syndrome) and acalculous cholecystitis. Trimethoprim–sulfamethoxazole provides an effective treatment for cyclosporiasis, and without therapeutic intervention, the disease may assume a remitting–relapsing course. C. cayetanensis is distributed in various parts of the world including countries of Central and South America, Asia, Africa, and Europe, and Australia as it can persist on food, in water and in the environment.1,15–25 The parasite has been isolated from sewage water, plants, and animal feces (e.g., chicken, ducks, dogs, rats, and mice),26–33 although the role of animals in the transmission of human cyclopsoriasis is unknown. In fact, as Eberhard et al.34 failed to find morphologically compatible oocysts in chicken, ducks, turkeys, pigeons, pigs, cattle, horses, goats, dogs, cats, and guineapigs, which lived in or near houses with human infection, the authors concluded that man is the only host for C. cayetanensis, and humans acquire the infection by way of human stool-contaminated food or water.12 Because C. cayetanensis is unsporulated and thus noninfectious when discharged in stool, it is incapable of direct fecal-oral infection. Food items associated with the disease outbreaks include raspberries, blackberries, fresh basil, fresh baby lettuce leaves, potato salad, etc. For example, human cyclosporiasis outbreaks that caused acute diarrhea in the United States and Canada between the years 1996 and 1998 were traced to the imported, unwashed raspberries.3,35 Other human cyclosporiasis outbreaks in the United States and Germany were linked to the consumption of fresh vegetables in salads and side-dishes.3,36–38 Given that coccidial oocysts are resistant to chlorine and most other chemical disinfectants, inadequately filtered drinking water as well as untreated or poorly treated water used for irrigation of fruits and vegetables may act as a vehicle of transmission for the parasite. Additionally, swimming pools were also found to harbor oocysts of C. cayetanensis. Cyclosporiasis appears to be a seasonal disease with nearly all (97.8%) of cases of cyclosporiasis occurring during the spring and summer months (April through July).

Cyclospora

Being an obligate intracellular pathogen, C. cayetanensis resides in epithelial cells of human intestines where it reproduces both asexually (via merogony) and sexually (via gametogony). The parasite is known to generate an electron-dense phospholipid membrane/myelin-like material in the intestine that appears to increase with length of the disease.39 It also induces other inflammatory changes in the infected areas of the upper small intestine (particularly the jejunum) and other parts of the body that are attributable to the diarrhea and clinical symptoms associated with cyclosporiasis in humans.

47.1.3  Diagnosis 47.1.3.1  Phenotypic Techniques Considering that C. cayetanensis typically produces cycles of diarrhea with anorexia, malaise, nausea, and cramping as well as a low-grade fever, which resemble those caused by Cryptosporidium, microsporidia, and Isospora belli,40 it is necessary to apply laboratory diagnostic techniques to differentiate C. cayetanensis from other parasitic and bacterial pathogens, to trace the source of an outbreak and to monitor vehicles of transmission.41 In general, identification of protozoa involves concentration, purification, detection, and viability assessment. For diagnosis of cyclosporosis in symptomatic or asymptomatic individuals, it can be achieved on the basis of the following criteria: (i) observation of C. cayetanensis oocysts in stool by microscopic examination, which are of characteristic size and shape (8–10 μm as measured with a microscope eyepiece graticule and spherical); (ii) recovery of oocysts in intestinal fluid or biopsy specimens; (iii) observation of two sporozoites each containing two sporocysts after in vitro oocyst sporulation; or (iv) detection of C. cayetanensis DNA by nucleic acid amplification. While the first three criteria are dependent on the application of phenotypic procedures that are often timeconsuming and imprecise, the fourth criterion relies on the measurement of genetic features that are intrinsically stable. Microscopic examimination provides a straightforward way to diagnose C. cayetanensis oocyst, although its efficiency can be variable since C. cayetanensis is relatively small, located intracellularly, and incapable of uptaking various stains. Therefore, phase contrast microscopy is often used to check for the spherical oocysts characterisitic of C. cayetanensis, modified acid-fast staining is performed to confirm acid-fast oocysts, and autofluorescence with UV lights is applied to identify oocysts (as C. cayetanensis oocysts autofluoresce dark blue under a 365-nm dichromatic filter, and autofluoresce mint green under a 450–490-nm dichromatic filter). Additionally, use of safranin can aid in the visualization of C. cayetanensis oocyst.42 For rapid screening of large numbers of samples, flow cytometry method can be employed.43 Frequently, centrifugation of a sample of feces in a sucrose solution (also known as sucrose or Sheather’s flotation technique) prior to mounting to a slide increases the sensitivity of microscopic detection of C. cayetanensis (to around 84%).

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The fact that oocysts sometimes give the appearance in wet mount slides as nonrefractile spheres under light microscope, and demonstrate variability in staining with a modified Ziehl–Nielsen stain and acid fast stain44 often complicates the direct micoscopic observation of C. cayetanensis oocyst in stool. In particular, it is known that the takeup of the acid fast stain by C. cayetanensis varies from nonstaining to full staining, in spite of the value of this stain for diagnosis of C. cayetanensis in stool specimens. This can lead to its misidentification. Attention should be also paid to the confustion caused by the co-presence of Cryptosporidium parvum in stool. Besides being smaller than C. cayetanensis, C. parvum appears red using modified acid fast staining and does not autofluorescence under UV light while C. cayetanensis does. This is especially so when dealing with environmental samples where presence of other substances tends to create additional difficuty for microscopic detection. To enhance its detection, it is useful to apply filtration, centrifugation, and flotation techniques to recover C. cayetanensis oocysts from food products and environmental specimens in which a relatively low number of organisms occur in contrast to stool containing about 102–104 oocysts per gram. Depending on the washing and detection procedures utilized, the rates of recovery from leafy vegetables and herbs ranged from 12 to 14%,45,46 they were higher from raspberries.45,47,48 The next step in the identification of C. cayetanensis involves in vitro sporulation, which is normally performed by subjecting the recovered oocysts to a 2-week incubation in 2.5% potassium dichromate solution.29,49 This not only helps ascertain the characteristic feature of C. cayetanensis oocysts (i.e., two sporocysts each containing two sporozoites), but also offers a means to determine the viability of organisms, which is vital for assessment of its risk associated with potentially contaminated water.50 Although conventional techniques based on histochemical staining and microscopy are still largely in use, they do not always provide a conclusive determination of C. cayetanensis species identity. While filtration, centrifugation, and flotation techniques are useful for recovery of C. cayetanensis oocysts from clinical, food, and environmental samples, they tend to be labor-intensive and show varied efficiency in dealing with different sample types. In vitro sporulation is undoubtedly valuable for confirmation of C. cayetanensis identity, but the length of time it takes to complete compromises its use in epidemic investigations where speedy identification is desired. 47.1.3.2  Molecular Techniques Molecular techniques such as PCR provide a more sensitive, specific and rapid alternative to conventional, phenotype-based methods for diagnosis of C. cayetanensis.51–55 Shortly after the description of C. cayetanensis as a human pathogen by Ortego et al.4 a nested PCR assay with primers (F1E-R2B, F3E-R4B) targeting 18S ribosomal DNA gene sequences of eukarya and C. cayetanensis, respectively, was devised by Relman et al.56 for C. cayetanensis identification. This method has been used successfully for sensitive

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detection of C. cayetanensis oocysts in human stool.47,48,57 To eliminate the cross-reaction of these primers with some Eimeria species (showing the identical size of a 294-bp amplicon), a restriction enzyme MnlI digestion step was incorporated into the assay (i.e., PCR-restriction fragment length polymorphism, or PCR-RFLP) that helped differentiate C. cayetanensis from Eimeria species.47 This modification extended the applicability of the assay to foodstuffs.37,47 and environmental waters.58 Following the identification of three new Cyclospora species: C. cercopitheci, C. colobi, and C. papionis from nonhuman primates in 1999 (see Table 47.1),59,60 it is apparent that the 18S rDNA regions from these species demonstrate a high similarity to that of C. cayetanensis, and the use of MnlI digestion in the PCR does not permit discrimination among the species. This lead to the development of another nested PCR assay by Shields and Olson,53 which empoys three new primers from 18S rDNA sequences from Cyclospora and Eimeria species: AO1-AR1 and AI2-AR1. After digestion with restriction enzyme AluI, C. cayetanensis can be distinguished from Eimeria and other Cyclospora species in water and soil samples without microscopic confirmation. In addition, focusing on the 18S rRNA gene of C. cayetanensis as well as those of nonhuman primate-derived Cyclospora spp, Chu et al.61 described a multiplex PCR for differentiation of primate-derived Cyclospora and Eimeria from human C. cayetanensis. Whereas primer pair CC719CRP999 amplifies a 298-bp fragment from the 18S rRNA gene of C. cayetanensis, primer pair PDCL661-CRP999 produces a 361-bp band from the 18S rRNA gene of nonhuman primate-derived Cyclospora spp, and primer pair ESSP841–CRP999 forms a 174-bp amplicon with the 18S rRNA gene of Eimeia spp. Without the need to digest the amplicons by restriction enzyme, this multiplex PCR enables rapid discrimination of Cyclospora and Eimeria in an efficient and precise manner. Further, Orlandi et al.62 took advantage of minute differences at the 18S SSU-rRNA gene amont Cyclospora spp. and designed a series of genus- and species-specific forward primers that distinguish sites of limited sequence heterogeneity in the same gene regions for differentiation between C. cayetanensis, nonhuman primate species of Cyclospora, and Eimeria species. Hussein63 also utilized a multilex PCR for detection of Cyclospora organisms in children suffering diarrhea. Moreover, Varma et al.43 reported a real-time TaqMan assay for C. cayetanensis detection that also targets the 18S rRNA gene sequences. Using the forward primer (CcayF1) and reverse primer (CcayR1) together with the probe (CcayP1), the authors showed that the assay was highly specific for C. cayetanensis as non-C. cayetanensis control organisms were unvariably negative. Hussein et al.64 described a RT-PCR for detection of C. cayetanensis in stool with a species-specific primer set and dual fluorescent labeled C. cayetanensis hybridization probe which originated from unique regions of 18S rRNA gene sequence. All symptomatic and asymptomatic patients were detected.

Molecular Detection of Foodborne Pathogens

In addition to 18S rDNA regions, the internal transcribed spacer region 1 (ITS1) of C. cayetanensis has been used for PCR identification of C. cayetanensis.65,66 The ITS genebased PCR assay by Adam et al.65 was useful for identification of different genotypes of C. cayetanensis while that by Olivier et al.66 facilitated differentiation of C. cayetanensis from C. papionis. However, considering enormous variation among C. cayetanensis oocysts in the same stool sample, this latter PCR assay may have limited value in the determination of its geographic source.66 Even though PCR technology provides a highly effective way to generate multiple copies of gene target with purified nucleic acids for subsequent detection and analysis, it is relatively inefficient in dealing with clinical, food, and environmental samples. This is due to the fact that polymerase enzymes used in the PCR process are susceptible to inhibition by substances that are commonly found in unprocessed samples. Notable polymerase-interfering compounds include heme, hemoglobulin, lactoferrin, immunoglobulin G, leukocyte DNA and polysaccharides in blood; phenolics, glycogen, calcium ions, fat, and other organic substances in foods; phenolics, humic acids and heavy metals in environmental samples. To reduce and eliminate inhibitory materials from these samples, it is critical that efficient sample preparation methods are utilized. Additionally, since food, water, and other environmental samples often contain low numbers of C. cayetanensis, inclusion of an enrichment protocol may be necessary to increase organism number and thus assay sensitivity. However this extra step creates delay in test turnover. Toward this goal, Orlandi and Lampel48 described an extraction-free, FTA filter-based protocol to prepare C. cayetanensis DNA templates for PCR identification of C. cayetanensis oocysts. Originally designed as matrix for blood storage, FTA filter (Whatman Inc., Newton, MA) incorporates denaturants, chelating agents and ­f ree-radical traps that lyse C. cayetanensis on contact and sequesters DNA within the matrix while other materials remain unbound. The adsorbed C. cayetanensis can then be used for DNA preparation. Adopting this procedure to the PCR, the authors detected three to ten oocysts directly applied to filters, and ten to 30 oocysts in complex mixed samples (e.g., 100 g of fresh raspberries) (which amounted to as few as 0.3 oocysts per gram of raspberries). Robertson et al.46 utilized lectin wheat germ aggluntinin-coated paramagnetic beads to improve recovery and purification of C. cayetanensis oocysts. With an affinity for β-linked polyN-acetyl hexosamine-containing polysaccharides, which constitute a component of C. cayetanensis oocyst walls, lectin aids the isolation of C. cayetanensis oocyst from other substances. This approach helped to clarify and concentrate the final sample volume, and make microscopic examination easier. Lalonde and Gajadhar 67 described a reliable DNA extraction for C. cayetanensis oocysts in water for PCR detection, with a limit of between one and ten oocysts.

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Cyclospora

47.2  Methods 47.2.1 Sample Preparation 47.2.1.1 Sample Processing Feces. Upon collection, fecal specimens are examined by differential interference contrast microscopy for the presence of Cyclospora oocysts of 8–10 µm in size, and oocysts can be further identified by UV epifluorescence microscopy. Cyclospora oocysts are then concentrated by using a discontinuous sucrose density purification method4, or by using a 10% formalin-ethyl acetate concentration in conjunction with Fecal Parasite Concentrators (Evergreen Scientific, Los Angeles, CA). For the latter method, fecal specimens (100–200 μl of packed debris) are washed twice with 1 ml of distilled water and pelleted by centrifugation. The washed debris is was then suspended in 1 ml of distilled water and extracted with 0.25 ml of ethyl acetate, vortexed for 20–30 sec, and centrifuged. After removing the upper, organic phase, any debris at the interface, and the lower aqueous phase, the sedimented debris is then washed twice with 1.0 ml of distilled water, suspended in 1 ml of distilled water and passed through a glass wool column. Foods. Fresh fruits and vegetables (100 g) are washed in 100 ml distilled water in a container, and agitated for 30 min at room temperature.37,40 The wash liquid is decanted and centrifuged at 1,870×g for 20 min at 4°C. The resulting sediment is then suspended in 5–10 ml of distilled water and applied to a Poly-Prep chromatography column (Bio-Rad, Hercules, CA) containing tightly packed glass wool presoaked with distilled water. The column is washed once with 5 ml of distilled water, and the eluent is centrifuged at 20,000×g for 15 min at 4°C. The resulting pellet is then suspended in 50–100 μl of distilled water. Water. Being in large volume, water samples are filtered using cartridge, hollow-fibre ultra-filters or capsule filters to capture oocysts. An alternative method of collection and concentration is flocculation that is not affected by turbidity. Namely, calcium chloride and sodium bicarbonate are added to a water sample and the pH is raised to 10.0 with sodium hydroxide. Acting as a coagulant, calcium carbonate increases the density and diameter of any particle in the water, which help its settling to the bottom of the container. The water is aspirated and the calcium carbonate is dissolved with a small volume of a mild acid (10% sulphamic acid).68 Resultant pellets are added with digestion buffer for genomic DNA extraction and or 2.5% potassium dichromate for microscopic analysis. 47.2.1.2 DNA Isolation (i) Phenol-chloroform extraction: The pellet is digested with proteinase K overnight, and centrifuged at 14,000×g for 1 min to remove debris. Phenol-chloroform is then used to remove proteins. The resultant aqueous layer is precipitated

with ethanol-NH4 acetate and washed with cold 70% ethanol. The purified DNA is dissovled in TE buffer. (ii) FastDNA Spin Prep Kit: Da Silva et al.69 used the FastDNA Spin Prep Kit for Cryptosporidium and Microsporidia genera in stool samples by the addition of polyvinylpyrrolidone (0.5% w/v) during the protein precipitation step, which helped removal of additional PCR inhibitors. Briefly, in 100 μl of packed pellet, 300 μl of digestion buffer and 50 mg of glass beads are added. After agitation in a bead beater, the crude DNA lysate is briefly centrifuged (~6,000×g for 30 sec), the supernatant is removed, and more buffer and detergent are added to the pellet. This is repeated for a total of four times, and the final centrifugation is at 14,000×g for 5 min. The crude lysates are pooled, and crude DNA is precipitated overnight in 5 M ammonium acetate (0.1× volumes) and ethanol (2.2 volumes) at −20°C. The crude DNA samples are then centrifuged (20 min at 14,000×g), and the resultant pellet is washed with 70% cold ethanol, air dried, and then resuspended in 500 μl of phosphate buffer for further purification. (iii) FTA filter purification: After spotting 25–50 μl of packed pellet (or 10–20 µl of stool) onto FTA filter and subsequent drying on a 56°C heating block, 6-mm FTA disks are punched out using an individual hole-punch, and placed in a 1.5-ml microcentrifuge tube. FTA disks are washed twice with 0.5 ml of FTA purification buffer (Life Technologies, Gaithersburg, Md.) for 2 min, twice with 0.5 ml of 10 mM Tris (pH 8.0) containing 0.1 mM EDTA for 2 min and again air-dried on a 56°C heating block. The washed filters are then used directly as the source of template in PCR.48

47.2.2  Detection Procedures 47.2.2.1  PCR-RFLP (i) PCR with F1E-R2B, F3E-R4B primers and RFLP with MnlI: Principle: A nested PCR protocol based on 18S rDNA gene was first developed by Relman et al.56 which was later modified by Orlandi’s group.48,61 This method has been widely applied for routine detection of C. cayetanensis. The first round PCR utilizes primers F1E (5′-TACCCAATGAAAACAGTTT-3′) and R2B (5′-CAGGAGAAGCCAAGGTAGG-3′), leading to a product of 636 bp; and the second round PCR uses primers F3E (5′-CCTTCCGCGCTTCGCTGCGT-3′) and R4B (5′-CGTCTTCAAACCCCCTACTG-3′), leading to a product of 294 bp from both Cyclospora spp. and Eimeria spp. Digestion of the second round PCR product with restriction enzyme Mnl I results in the formation of 140, 106, and 48 bp fragments for Cyclospora and 126, 106, and 62 bp fragments for Eimeria.61

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Procedure: (1) Prepare the first round PCR mixture (100 µl) containg Hot-StarTaq Master Mix (Qiagen, Valencia, CA), containing 2.0 mM MgCl2, and 200 µM of each dNTP, 0.2 µM of F1E and R2B primers, and 1–5 µl of purified DNA. (2) Conduct the first round PCR amplification with one cycle of 95°C for 15 min (for activation of the HotStarTaq DNA polymerase); 35 cycles of 94°C for 30 sec, 53°C for 30 sec, and 72°C for 90 sec; one cycle of 72°C for 10 min followed by soaking at 4°C. Run 10 µl of the PCR product on 1% agarose gel to verify the successful amplification of the 636 bp fragment. (3) Prepare the second round PCR mixture (50 µl) with HotStarTaq Master Mix, 2.0 mM MgCl2, 200 µM of each dNTP, and 0.2 µM each of primers F3E and R4B primers, and 1–5 µl of the first round PCR product as the template. (4) Conduct for the secondary nested PCR amplification with one cycle of 95°C for 15 min (for activation of the HotStarTaq DNA polymerase); 35 cycles of 94°C for 30 sec, 60°C for 30 sec, and 72°C for 90 sec; one cycle of 72°C for 10 min followed by soaking at 4°C. (5) Load 10 µl PCR amplicon on a 1.5% agarose gel and stain with 0.2 µg/ml of ethidium bromide. Visualize the the specific amplicon of 294 bp under UV transillumination. (6) Add 2.5 µl of 10× reaction buffer and 2 U Mnl I (New England Biolabs, Beverly, MA) to 20 µl of the secondary PCR amplicon (total volume 25 µl), and incubate for 2 h at 37°C. (7) Separate the digests on a 5% Nusieve 3:1 agarose gel (Biowittaker Molecular Applications, Rockland, ME) containing 0.2 µg/ml of ethidium bromide and visualize under UV transillumination. Note: Although this routinely used nested PCR-RFLP protocol is capable of detecting and distinguishing Cyclospora from Eimeria spp, it is unable to differentiate between C. cayetanensis and non-human primate-derived Cyclospora spp. (ii) PCR with AO1-AR1, AI2-AR1 primers and RFLP with AluI: Principle: While the PCR-RFLP protocol with F1E-R2B, F3E-R4B primers and RFLP with MnlI56 is capable to identify C. cayetanensis oocysts in stool samples, it requires microscopic confirmation to produce reliable results for environmental samples, due to the fact that C. cayetanensis, C. cercopitheci, C. colobi, and C. papionis all show a high degree of homology in the inner (CYCF3E and CYCR4B) PCR products that are not distinguished with MnlI digestion. Therefore, Shields and Olson52 designed three new primers from 18S rDNA sequences from Cyclospora and Eimeria species: AO1 (5′-ATAACGAACGAGACCTTAGCCT-3′), AR1 (5′-AA-GGATGCAAAAGTCGTAACAC-3′), AI2 (5′-CAGGTCT-GGGTAATCTTTTGAG-3′). The first round

Molecular Detection of Foodborne Pathogens

PCR is performed with AO1-AR1, and the second round PCR with AI2-AR1. This is followed by RFLP with restriction enzyme AluI. The method allows differentiation of C. cayetanensis from Eimeria and other Cyclospora species in water and soil samples without microscopic confirmation. Procedure: (1) Prepare first round PCR mixture (50 μl) containing 50 mM KCl, 10 mM Tris-HCl pH 8.3, 200 μg of nonfat dry milk, 200 μM each deoxynucleoside triphosphate, 2.0 mM MgCl2, 0.4 μM each of AO1 and AR1 primers, 1 U of AmpliTaq polymerase, and 5 μl of template. (2) Conduct the first round PCR amplification for one cycle of 94°C for 5 min; 45 cycles of 94°C for 30 sec, 59.5°C for 30 sec, and 72°C for 90 sec; and one cycle of 72°C for 9 min. (3) Prepare the second round PCR mixture (50 μl) containing 50 mM KCl, 10 mM Tris-HCl pH 8.3, 200 μM each deoxynucleoside triphosphate, 2.0 mM MgCl2, 0.4 μM each of AI2 and AR1 primers, 1 U of AmpliTaq polymerase, and 2.5 μl of the first round PCR product. Conduct the second round PCR amplification as above. (4) Load 10 μl PCR product on 1.75% agarose gels supplemented with 0.5 μg of ethidium bromide per ml at constant current in Tris-borate-EDTA buffer. Include three molecular size markers: 25- and 50-bp ladders (Promega) and DNA weight marker VIII (a mixture of pUCBM21 DNA cleaved with HpaII and pUCBM21 DNA cleaved with DraI and HindIII; Boehringer Mannheim, Indianapolis, IN). Visualize the product by UV transillumination. (5) Digest the positive PCR products from the second reaction (10 μl) with 1 U of AluI (New England Biolabs) at 37°C for 2 h. Separate the fagments, visualize, and photograph as described above. Note: The expected second round PCR product from Cyclospora species using AI2 and AR1 primers is 251 bp, which shows four bands at 98, 88, and 50 bp and a faint band at 15 bp for C. cayetanensis after AluI digestion. However, the second round PCR product becomes 153–159 bp and/or 94–98 bp for Eimeria and C. papionis and three bands for Isospora robini and the other two Cyclospora species after AluI digestion. 47.2.2.2  Multiplex PCR Principle: A multiplex PCR was described by Chu et al.61 to distinguish the presence of primate-derived Cyclospora and Eimeria from human C. cayetanensis. Using the first round PCR product of 636 bp (which is generated with ­primers F1E: 5’-TACCCAATGAAAACAGTTT-3′ and R2B: 5′-CAGGAGAAGCCAAGGTAGG-3′) as tmplate, the multipex PCR is conducted with common reverse primer CRP999 (5′-CGTCTTCAAACCCCCTACTGTCG-3′), species-specific

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forward primers CC719 (5′-GTAGCCTTCCGCGCTTCG-3′); PDCL661 (5′-CTGTCGTGGTCATCTGTCCGC-3′), and ESSP841 (5′-GTTCTATTTTGTTGGTTTCTAGGACC A-3′). While primer pair CC719-CRP999 generates a 298-bp amplicon from the 18S rRNA gene of C. cayetanensis, primer pair PDCL661-CRP999 generates a 361-bp amplicon from the 18S rRNA gene of nonhuman primate-derived Cyclospora spp, and primer pair ESSP841–CRP999 generates a 174-bp amplicon from the 18S rRNA gene of Eimeia spp. Procedure: (1) Prepare multiplex PCR mixture (50 µl) containing HotStarTaq Master Mixture, 2 mM MgCl2, 0.2 µM of CRP999, CC719, PDCL661, and ESSP841 primers and 1–5 µl of PCR product generated with F1E: and R2B (as shown in Section 47.2.2.1). (2) Perform PCR amplification with one cycle of 95°C for 15 min; 25 cycles of 94°C for 15 sec and 66°C for 15 sec. (3) Separate the PCR products on a 1.5% agarose gel with 0.2 µg/m of ethidium bromide and visualize on a UV transilluminator. Note: The primer pair CC719-CRP999 generates an amplicon of 298 bp only with C. cayetanensis; the primer pair PDCL661-CRP999 generates an amplicon of 361 bp with other Cyclospora spp. (e.g., C. cercopitheci); and the primer pair ESSP841-CRP999 generates an amplicon of 174 bp with Emeria spp. (e.g., E. tenella).61,62 47.2.2.3 Real-time PCR Principle: A real-time C. cayetanensis-specific TaqMan assay targeting 18S rRNA gene sequences was developed by Varma et al.43 The forward primer (CcayF1) is located at nucleotide positionsof1671–1692(5′-TGAACTCATTGGACTGACCAGC-3′), the reverse primer (CcayR1) at 1753–1731 (5′-ACTTTTG CATCCTTTAGAGGGCT-3′) and the probe (CcayP1) at 1698– 1721 (5′-FAM-TTCGCGGAGCTGGTCGGAAAGTTG-TAMRA-3′), based on the C. cayetanensis sequence in GenBank accession no. AF111183. While the forward and reverse primers are synthesized unlabeled, the probe is synthesized with dual labels having a reporter fluorescent dye [6-carboxyfluorescein (6-FAM)] on the 5′-end and a quencher fluorescent dye [6-carboxytetramethyl-rhodamine (TAMRA)] on the 3′-end. The 3′-end of the probe is also phosphorylated to prevent probe extension. Procedure: (1) Prepare real-time PCR mixture (25 μl) in 0.5-ml MicroAmp optical PCR tubes (Applied Biosystems) containing 5 μl of extracted C. cayetanensis DNA (diluted as described), 2.4 μl of a 450 nM solution of the dual-labeled C. cayetanensis-specific hybridization probe (CcayP1), 2.4 μl of a 5.2 μM solution of the forward primer (CcayF1), 2.4 μl of a 5.2 μM solution of the reverse primer (CcayR1), 0.9 μl of a

2 mg/ml bovine serum albumin Fraction V (SigmaAldrich), and 11.9 μl of the of the 2× TaqMan Universal PCR Master Mix [AmpliTaq Gold DNA polymerase, Amperase UNG, dNTPs, passive reference 6-carboxy-X-rhodamine (ROX), MgCl2, and buffer components in amounts undisclosed by the manufacturer] (Applied Biosystems). Insert the optical tubes into a 96-well tray/retainer set and seal. Centrifuge the tray is for 5 min at 500×g before inserting into an ABI-Prism 7700 Sequence Detector (Applied Biosystems). Five replicates are prepared for each sample. (2) Conduct PCR amplification for one cycle of 50°C for 2 min; one cycle of 95°C for 10 min; 40 cycles of 95°C for 0.25 min and 60°C for 1 min. Note: The forward primer shows a high degree of specificity for the C. cayetanensis ribosomal DNA gene, while the reverse primer is broadly specific for a variety of protists, the hybridization probe is also specific to several Cyclospora spp. The combination of forward primer and probe specificity results in a highly specific assay, and with a CT value of at least 40, no positive results is obtained with non-C. cayetanensis control organisms tested.43

47.3 Conclusions and Future Perspectives The genus Cyclospora consists of at least 18 known species of protozoa sharing morphological, biological and molecular similarities with Cryptosporidium, Isospora and Sarcocystis.6 Being obligate intracellular parasites, 14 of the 18 Cyclospora species are found in insects and reptiles, three (C. colobi, C. cercopitheci, and C. papionis) are infective to nonhuman primates, while one (C. cayetanensis) causes cyclosporiasis in humans after gaining entry into the host via contaminated food and water (Table 47.1). Given that the clinical manifestations of human cyclosporiasis are nonspecific (e.g., diarrhea and general illnesses), which are indifferent from those of Cryptosporidium, Isospora, Sarcocystis and other bacterial and viral infections, correct identification of C. cayetanensis is essential for selecting appropriate chemotherapy and effective control measures. Conventional techniques for identification of C. cayetanensis are largely based on the morphological features of this parasite. Specifically, the ocysts (8×10 μm) of C. cayetanensis are larger than those of Cryptosporidium (5.2×4.9 μm), but smaller than those of Isospora belli (30×12 μm in diameter) and Sarcocystis (30×15 μm). In addition, C. cayetanensis oocysts comprise two sporocysts each containing two sporozoites, Cryptosporidium oocysts have no sporocyst but four sporozoites, Isospora and Sarcocystis oocysts all possess two sporocysts each containing four sporozoites. Therefore, through a combination of microscopic examination and in vitro sporulation, it is possible to differentiate Cyclospora from Cryptosporidium, Isospora and Sarcocystis. Use of

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immunological assays such as ELISA adds another dimension on the phenotypic characterization of Cyclospora spp. However, besides being time-consuming and lacking precision, these phenotypic techniques do not allow precise discrimination among Cyclospora species. The development and application of nucleic acid amplification technologies such as PCR during the past decades have facilitated improved laboratory identification and diagnosis of C. cayetanensis. By employing primers derived from 18S rRNA genes, intergenic spacer (ITS) and other gene regions in various PCR formats (e.g., standard PCR, PCR-RFLP, multiplex PCR, real-time PCR and reversetranscription PCR), C. cayetanensis can be rapidly, precisely and sensitively distinguished from closely related C. colobi, C. cercopitheci and C. papionis as well as Eimeria species. These “state-of-art” molecular detection procedures offer a valuable tool in the laboratory diagnosis of C. cayetanensis and epidemiological investigations of human cyclosporiasis outbreaks, contributing to improved control and prevention of this important foodborne disease. Nonetheless, while the PCR assays generally give remarkably reliable performance with purified nucleic acids as template, their efficiency is somewhat compromised when clinical, food and environmental samples are used directly. This is attributable to the fact that polymerase enzymes used in PCR are susceptible to interference to inhibitory substances that are abundantly present in unprocessed clinical, food and environmental samples. Although a range of prior sample processing procedures (e.g., filtration, centrifugation, and concentration) together with nucleic acid isolation protocols are useful to reduce and eliminate the interfering materials from the samples, there is a lack of consensus with regard to the best approach to this issue. This is especially important in cases where only low numbers of C. cayetanensis oocysts exist in the samples (e.g., environmental water and food samples). Thus, there is a continuing need to develop, evaluate and optimize sample processing procedures to enhance recovery of C. cayetanensis oocysts from environmental water and food samples. Considering that food samples alone come in vast variety and complexity, the challenge is enormous in finding and designing efficient processing procedures that are suited to distinct sample types. The availability of these highly efficient, individualized sample prepareation procedures will contribute to the improve performance of PCR assays for direct detection and identification of C. cayetanensis from clinical, food and environmental samples. Additionally, there is scope for further improvement of the applicability of the molecular detection techniques in field situations or in places where access to expensive equipment is resticted and where rapid and precise diagnostic techiques for C. cayetanensis are most needed. Moreover, future development of a reliable and effective animal model for C. cayetanensis infection will lead to new insights on the pathogenic mechanisms of cyclosporiasis.70

Molecular Detection of Foodborne Pathogens

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44. Abou el Naga, I.F. Studies on a newly emerging protozoal pathogen: Cyclospora cayetanensis. J. Egypt. Soc. Parasitol., 29, 575, 1999. 45. Ortega, Y.R. et al. Pathologic and clinical findings in patients with cyclosporiasis and a description of intracellular parasite life-cycle stages. J. Infect. Dis., 176, 1584, 1997. 46. Robertson, L.J., Gjerde, B. and Campbell, A.T. Isolation of Cyclospora oocysts from fruits and vegetables using lectincoated paramagnetic beads. J. Food Prot., 63, 1410, 2000. 47. Jinneman, K.C. et al. Template preparation for PCR and RFLP of amplification products for the detection and identification of Cyclospora sp. and Eimeria spp. oocysts directly from raspberries. J. Food Prot., 61, 1497, 1998. 48. Orlandi, P.A. and Lampel, K.A. Extraction-free, filter-based template preparation for rapid and sensitive PCR detection of pathogenic parasitic protozoa. J. Clin. Microbiol., 38, 2271, 2000. 49. Sturbaum, G.D. et al. Species-specific, nested PCRrestriction fragment length polymorphism detection of single Cryptosporidium parvum oocysts. Appl. Environ. Microbiol., 67, 2665, 2001. 50. Dalton, C. et al. Viability of Giardia intestinalis cysts and viability and sporulation state of Cyclospora cayetanensis oocysts determined by electrorotation. Appl. Environ. Microbiol., 67, 586, 2001. 51. Quintero-Betancourt, W., Peele, E.R. and Rose, J.B. Cryptosporidium parvum and Cyclospora cayetanensis: a review of laboratory methods for detection of these waterborne parasites. J. Microbiol. Methods, 49, 209, 2002. 52. Shields, J.M. and Olson, B.H. PCR-restriction fragment length polymorphism method for detection of Cyclospora cayetanensis in environmental waters without microscopic confirmation. Appl. Environ. Microbiol., 69, 4662, 2003. 53. Steele, M., Unger, S. and Odumeru, J. Sensitivity of PCR detection of Cyclospora cayetanensis in raspberries, basil, and mesclun lettuce. J. Microbiol. Methods, 54, 277, 2003. 54. Veiweij, J.J. et al. Detection of Cyclospora cayetanensis in travellers returning from the tropics and subtropics using microscopy and real-time PCR. Int. J. Med. Microbiol., 293, 199, 2003. 55. Mundaca et al. Use of PCR to improve diagnostic yield in an outbreak of cyclosporiasis in Lima, Peru. Trans. R. Soc. Trop. Med. Hyg., 102, 712, 2008. 56. Relman, D.A. et al. Molecular phylogenetic analysis of Cyclospora, the human intestinal pathogen, suggests that it is closely related to Eimeria species. J. Infect. Dis., 173, 440, 1996. 57. Jinneman, K.C. et al. An oligonucleotide-ligation assay for the differentiation between Cyclospora and Eimeria spp. polymerase chain reaction amplification products. J. Food Prot., 62, 682, 1999. 58. Sturbaum, G.D. et al. Detection of Cyclospora cayetanensis in wastewater. Appl. Environ. Microbiol., 64, 2284, 1998. 59. Eberhard, M.L et al. Morphologic and molecular characterization of new Cyclospora species from Ethiopian monkeys: C. cercopitheci sp. n., C. colobi sp. n., and C. papionis sp. n. Energ. Inf. Dis., 5, 651, 1999. 60. Lopez, A.S. et al. Epidemiology of Cyclospora cayetanensis and other intestinal parasites in a community in Haiti. J. Clin. Microbiol., 41, 2047, 2003. 61. Chu, D.M. et al. Detection of Cyclospora cayetanensis in animal fecal isolates from Nepal using an FTA filter-base polymerase chain reaction method. Am. J. Trop. Med. Hyg., 71, 373, 2004.

676 62. Orlandi, P.A. et al. Targeting single-nucleotide polymorphisms in the 18S rRNA gene to differentiate Cyclospora species from Eimeria species by multiplex PCR. Appl. Environ. Microbiol., 69, 4806, 2003. 63. Hussein, E.M. Molecular identification of Cyclospora spp. using multiplex PCR from diarrheic children compared to others conventional methods. J. Egypt .Soc. Parasitol., 37, 585, 2007. 64. Hussein, E.M. et al. Real-time PCR and flow cytometry in detection of Cyclospora oocysts in fecal samples of symptomatic and asymptomatic pediatrics patients. J. Egypt. Soc. Parasitol., 37, 151, 2007. 65. Adam, R.D. et al. Intervening transcribed spacer region 1 variability in Cyclospora cayetanensis. J. Clin. Microbiol., 38, 2339, 2000.

Molecular Detection of Foodborne Pathogens 66. Olivier, C. et al. Sequence variability in the first internal transcribed spacer region within and among Cyclospora species is consistent with polyparasitism. Int. J. Parasitol., 31, 1475, 2001. 67. Lalonde, L.F. and Gajadhar, A.A. Highly sensitive and specific PCR assay for reliable detection of Cyclospora cayetanensis oocysts. Appl. Environ. Microbiol., 74, 4354, 2008. 68. Vesey, G. et al. A new method for the concentration of Cryptosporidium oocysts from water. J. Appl. Bacteriol., 75, 82, 1993. 69. da Silva, A.J. et al. Fast and reliable extraction of protozoan parasite DNA from fecal specimens. Mol. Diagn., 4, 57, 1999. 70. Eberhard, M.L. et al. Attempts to establish experimental Cyclospora cayetanensis infection in laboratory animals. J. Parasitol., 86, 577, 2000.

48 Entamoeba Damien Stark

St. Vincent’s Hospital

John Ellis

University of Technology Sydney

Contents 48.1 Introduction.................................................................................................................................................................... 677 48.1.1 History, Classification, and Morphology.......................................................................................................... 677 48.1.2 Entamoeba Species that Infect Humans........................................................................................................... 678 48.1.3 Life Cycle and Epidemiology........................................................................................................................... 678 48.1.4 Pathogenesis, Clinical Presentation, and Treatment......................................................................................... 679 48.1.4.1 Pathogenesis..................................................................................................................................... 679 48.1.4.2 Clinical Presentation . ..................................................................................................................... 680 48.1.4.3 Treatment ........................................................................................................................................ 680 48.1.5 Conventional and Molecular Diagnosis............................................................................................................ 681 48.1.5.1 Microscopy...................................................................................................................................... 681 48.1.5.2 Culture............................................................................................................................................. 681 48.1.5.3 Isoenzyme Analysis......................................................................................................................... 681 48.1.5.4 Antibody Detection.......................................................................................................................... 681 48.1.5.5 Antigen Detection............................................................................................................................ 681 48.1.5.6 Molecular Diagnosis........................................................................................................................ 682 48.2 Methods.......................................................................................................................................................................... 684 48.2.1 Reagents and Equipment ................................................................................................................................. 684 48.2.2 Sample Collection and Preparation.................................................................................................................. 686 48.2.3 Detection Procedure......................................................................................................................................... 686 48.3 Conclusions and Further Perspectives............................................................................................................................ 687 References.................................................................................................................................................................................. 687

48.1  Introduction Entamoeba histolytica is an intestinal protozoan parasite and the causative agent of human amebiasis. It remains a significant cause of morbidity and mortality worldwide. It is estimated that one-tenth of the world’s population is infected with E. histolytica and this infection results in up to 100,000 deaths each year.1 E. histolytica has a worldwide distribution but is more commonly found in developing countries, and is more prevalent in the tropics and sub tropics.2

48.1.1 History, Classification, and Morphology E. histolytica (phylum Sarcomastigophora, family Endamoebidae) is defined as a unicellular eukaryote whose life cycle contains two main stages: trophozoites (diameter 20–40 µm) with a single nucleus and cysts (diameter 10–16 µm) with four nuclei when mature; the immature cyst has a single nucleus often with a small central karyosome.3

The trophozoite form of E. histolytica was first described in 1875 by Fedor Losch, from dysenteric stools and colonic ulcerations from a Russian farmer with a fatal case of dysentery.4 Although Losch was able to induce dysentery in a dog that was inoculated rectally with the patients stools, he failed to recognize the causal relationship between the organism and disease. It was not until 1891 that clinical and pathological evidence for the association of E. histolytica with dysentery and liver abscesses was described by Councilman and LaFleur.5 In 1903 Quincke and Roos described the cyst form of the organism and in 1903, Schaudin named it E. histolytica and differentiated it from other Entamoeba species.6 Leonard Rogers designated emetine as the first effective treatment for amebiasis in 1912.6 In 1913 Walker and Sellards conducted controlled experiments with E. histolytica on prisoners in the Philippines.7 They found that transmission occurred via cysts, not trophozoites, asymptomatic carriers acted as reservoirs and were responsible for transmission, there were differences between individuals’ disease risk and there were 677

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differences in organism strain virulence.7 In 1925, Brumpt formulated the theory of the existence of two distinct but morphologically identical species, namely, the pathogenic invasive form E. histolytica and the nonpathogenic, noninvasive form E. dispar.6 His hypothesis was dismissed at the time and gained little support until isoenzyme typing differentiated pathogenic and nonpathogenic strains.8 In 1993 Clark and Diamond, on the basis of molecular, immunological, and biochemical analysis re-described the protozoa, retaining the name E. histolytica for the pathogenic invasive parasite, and renaming the noninvasive parasite E. dispar.9–10 The morphological characteristics of E. histolytica can be seen in Figure 48.1. E. histolytica trophozoites can vary in size from 10 to 50 µm, with most generally measuring between 15 and 20 µm. The trophozoites are unnucleated and may often exhibit pseudopoding motility which is often described as sporadic, rapid, and unidirectional. The cytoplasm of the trophozoite is finely granular and may contain ingested material. The nuclear structure consists of a centrally located, small, compact karyosome with evenly arranged peripheral chromatin present that may have a beaded appearance.11 Cysts of the organism range in size from 10 to 20 µm and may contain Cysts

Entamoeba histolytica

Trophozoites

one, two or four nuclei. Inclusions may be present in the cysts such as chromatoidal bars and a glycogen mass.11 The nuclear structure is similar to what is found in trophozoites. The morphology is often difficult to see in unstained preparations. When the organism is examined on a permanent stained smear the morphological characteristics are readily seen.

48.1.2  Entamoeba Species that Infect Humans There are many described species in the Genus Entamoeba several of which may infect humans, including those that reside in the intestinal lumen: E. histolytica, E. dispar, E. moshkovskii, E. coli, and E. hartmanni. There have also been reports of E. polecki and E. chattoni infections in humans though they are normally associated with pigs and nonhuman primates, respectively.12 E. histolytica is the only species pathogenic to humans; all the others are considered nonpathogenic.13 E. histolytica is morphologically identical to E. dispar and E. moshkovskii. Microscopy is unable to differentiate these three indistinguishable species.13 Until recently E. moshkovskii was considered to be a mainly a free living ameba, with environmental isolates recovered from sewage, fresh brackish water and salt water sediments.14 Reports on E. moshkovskii in human specimens have recently emerged from around the world.13,15–18 The existence of these morphologically indistinguishable species of Entamoeba highlights the need to differentiate the three species to determine clinically if the patient is infected with the pathogenic E. histolytica or the nonpathogenic E. dispar/E. moshkovskii. In a clinical setting a patient infected with the nonpathogenic E. dispar/E. moshkovskii could be treated unnecessarily with anti-amebic chemotherapy.

48.1.3 Life Cycle and Epidemiology

Figure 48.1  Cysts and trophozoitesof Entamoeba histolytica permanently stained using a modified iron-hematoxylin stain at 1000× magnification.

The life cycle is seen in Figure 48.2. The life cycle was first described by Dobell19 and unlike many protozoan parasites has a simple life cycle, comprising of trophozoites, precyst, cyst, metacyst, and metacystic trophozoites stages. The cyst form is the infective stage for humans and can survive in contaminated food and water.6 Ingestion of food and drink contaminated with E. histolytica cysts from human feces and direct fecal-oral contact are the most common means of infection. Infection begins with the ingestion of infective cysts. After cysts are ingested they survive the acid of the stomach and travel to the small intestine. Within the terminal ileum or colon, where the pH becomes neutral or slightly alkaline, the encysted organisms develop (during excystation) into four separate metacystic trophozoites which then mature into normal trophozoites that establish infections in the large intestine. Cyst formation occurs in the intestinal tract, and infective cysts are passed in the feces. Humans, and some nonhuman primates, are the only natural hosts.1 E. histolytica infections occur worldwide but are more prevalent in the tropics.6 The prevalence of infection varies from area to area as does the severity of disease from patient to patient. It has been suggested that 12% of the world’s

679

Entamoeba

5 1 5

5

5

4

2

3

(1) Cysts are ingested through contaminated food and water. (2) Cysts pass through the digestive system to the small intestine where excystation occurs, releasing trophozoites. (3) Trophozoites travel to the large intestine where they multiply by binary fission and colonize the intestinal lumen (nonvasive infection). Encystation occurs. (4) Cysts and trophozoites are passed in the feces to the external environment. Trophozoites degrade rapidly, while cysts can persist for weeks in the external environment. (5) Trophozoites may invade the intestinal mucosa and travel through the bloodstream to the brain, lungs liver and heart resulting in the formation of amebic abcesses (invasive infection). Amoebic abcesses most commonly occur in the liver.

Figure 48.2  Life cycle of Entamoeba histolytica.

population is infected with this organism.20 The epidemiology of E. histolytica remains uncertain as most of the existing data were obtained using methods incapable of distinguishing the three morphologically identical species. Most studies that have investigated the prevalence of E. histolytica and E. dispar have not considered the presence of E. ­moshkovskii. The worldwide prevalence of E. histolytica and E. dispar as separate species is not well defined and the prevalence of E. moshkovskii is unknown. E. dispar appears to be much more common than E. histolytica with most of the 500 million people infected with E. histolytica/E. dispar/E. ­moshkovskii carrying E. dispar.21 Most morbidity and mortality due to amebiasis occur in developing regions such as Central and South America, Africa, and the Indian subcontinent.22 In developed countries most infections are due to E. dispar. High risk groups for E. histolytica infection include immigrants from or travelers to areas of endemicity, men who have sex with men and institutionalized populations.18,23

48.1.4 Pathogenesis, Clinical Presentation, and Treatment 48.1.4.1  Pathogenesis E. histolytica is unique among the intestinal amebae as it has the ability to invade host tissue. The factors that control the

invasiveness of E. histolytica are incompletely understood. The main key to the pathogenesis of this organism is its ability to directly lyse host cells and cause tissue destruction. The mechanisms of pathogenicity include adherence, extracellular cytolysis and phagocytosis.24–26 Host cell leukocytes also play a major role in pathogenesis along with numerous other parasite factors.6,27 Disease begins with the adherence of E. histolytica to colonic epithelial cells. A number of E. histolytica adherence receptors have been identified, but the galactose-specific lectin receptor is thought to be responsible for mediating the attachment to colonic mucus and colonic epithelial cells.28 Human colonic epithelial cells and mucus contain a large number of N-acetylgalactosamine residues.29 Mammalian cells without N-terminal galactose or N-acetylgalactosamine residues are resistant to adherence by amebic trophozoites and to killing by E. histolytica, consistent with a physiological role for the lectin in adhesion.30 The cytolytic capabilities of E. histolytica have been demonstrated to be caused by amoebapores (a family of small peptides capable of forming pores in lipid bilayers). The pore forming proteins are transferred from the trophozoites to the target cell, causing disruption of the transmembrane gradient and subsequent lysis.31 Cysteine proteinases are also important virulence factors and are directly implicated in invasion

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and inflammation of the gut. These proteinases are major proteolytic enzymes and can degrade cellular attachment and matrix proteins such as collagen, laminin, and fibronectin. There is a direct correlation between the amount of proteinase activity and the pathogenicity of E. histolytica.32 The host’s cellular and humoral immune responses to E. histolytica infection also play a role in the pathogenesis of the organism. As invasive strains of E. histolytica have been shown to be resistant to complement-mediated lysis, cell mediated immunity represents the major human host defense.6 Intestinal epithelial cells produce interleukin-1B, interleukin-8, and cyclo-oxygenase (COX)-2 in response to E. histolytica.33 These mediators attract neutophils and macrophages. This inflammation can contribute to tissue damage early in disease. 48.1.4.2  Clinical Presentation Clinical syndromes associated with amebiasis vary widely with the host and the organism both playing a role. Syndromes range from asymptomatic infection to disseminated fatal disease. Asymptomatic infection. Most human E. histolytica infections are asymptomatic. It is estimated that 90% of those infected will not go on to develop disease.34 In such individuals the organism acts as a harmless commensal. Many asymptomatic individuals never become symptomatic and usually only excrete cysts in the stool for a short period, and often clear the infection without any signs of disease.35 This pattern is found in patients infected with pathogenic or nonpathogenic strains.10 However asymptomatic colonization with E. histolytica, if left untreated, can lead to disease and it is estimated that 4–10% of asymptomatic individuals may develop gastrointestinal or invasive disease.36 Symptomatic infection. Gastrointestinal. Approxi­ mately 10% of individuals infected with E. histolytica will develop clinical symptoms, with intestinal or extaintestinal involvement.34 Patients with noninvasive disease may exhibit nonspecific gastrointestinal symptoms including abdominal pain and diarrhea.6 The symptoms associated with invasive gastrointestinal amebiasis are usually also nonspecific. Patients with amebic colitis present with bloody diarrhea with abdominal pain and tenderness.37 The onset is usually gradual, over weeks, with abdominal pain, diarrhea, dysentery, and weight loss depending on severity of disease. When amebic colitis is diagnosed it is usually associated with ulceration of the colon. Perforation of the colon may occur in over 20% of patients with acute amebic.38 Perforations occur most commonly in the cecum and may result in peritonitis.39 Occasionally patients may develop a fulminant amebic colitis, with profuse bloody diarrhea, fever, leucocytosis, dehydration, and abdominal pain, often with peritoneal signs and extensive involvement of the colon.40 Complications of intestinal amebiasis include toxic megacolon, ameboma,41 cutaneous amebiasis42 and rectovaginal fistulae.43 Extraintestinal. Extraintestinal disease is the result of previous intestinal infection. Dissemination of E. histolytica

Molecular Detection of Foodborne Pathogens

infection to extraintestinal sites most frequently involves the liver and to a lesser extent the lungs and rarely pericardium, brain, and skin. Invasive amebiasis is associated with significant morbidity and mortality. Amoebic liver abcesses (ALA) is caused by the hematogenous spread of the invasive trophozoites from the colon, which reaches the liver via the portal vein.6 This explains the frequent occurrence of abscesses in the right hepatic lobe, which receives most of the blood draining from the cecum and ascending colon.44 ALA can occur concurrently with colitis, often there is intestinal infection and stool microscopy is usually negative for trophozoites, but more often they have no gastrointestinal symptoms. Onset may be acute or sub acute. Symptoms of upper right abdominal pain, fever, and substantial hepatic tenderness are commonly seen.34 Hematogeneous spread to other sites including the brain, lung, pericardium, and other sites is possible, with E. histolytica having been reported in almost every soft organ and tissue of the body. Amebiasis of the lungs may develop when hepatic abscesses rupture through the diaphragm, along with originating as a secondary site from the original intestinal infection. Rupture of ALA into the peritoneum or pericardium may occur.34 Amoebic brain abscesses are rare and usually arise from hepatic or pulmonary involvement. Sudden onset of symptoms and rapid progression to death has been reported in a majority of patients with amebic brain abscesses.45 Other ectopic sites include the urinary tract (kidney, ureters, urinary bladder, urethra), genital involvement (penis, clitoris, rectovaginal fistulaes, perianal region) and cutaneous lesions have all been described as case reports.46–48 Amebiasis cutis involves the skin and usually develops as a perianal extension of amebic colitis or on the abdominal wall due to rupture of intestinal or hepatic lesions. It may also occur as a venereal infection.49 48.1.4.3 Treatment Treatment of amebiasis is divided into two categories; treatment with luminal amebicides and treatment with tissue amebicides.34 Lumen agents used in the treatment of amebic infections include Diloxanide, Paromomycin, and Iodoquinol. These agents act on the organism in the intestinal lumen and are not effective against organisms in tissue.6 Tissue amebicides include the nitroimidazole derivatives (metronidazole, tinidazole, ornidazole), chloroquine or dehydroemetine. These agents are effective in the treatment of invasive amebiasis but are ineffective in the treatment of the organisms in the intestinal lumen.6 Asymptomatic carriers of E. histolytica should be treated with a luminal agent to not only minimize the spread of disease but to also prevent the risk of developing invasive disease.50 In patients with invasive disease a tissue amebicide should be used in conjunction with a luminal agent to eradicate the organism.6 The management of ALA is unique and can be cured without drainage, and even with one dose of metronidazole.51 The role of surgery is generally limited to patients with complications of invasive disease, as often cure

Entamoeba

can be achieved with medical therapy alone.52 All treated patients should be followed up clinically using appropriate laboratory tests.

48.1.5 Conventional and Molecular Diagnosis 48.1.5.1  Microscopy Numerous diagnostic techniques have been used for the diagnosis of amebic infection. Traditional diagnosis of amebiasis relies on conventional microscopy of wet preparations, concentrations, and permanently stained smears for the identification of E. histolytica in feces. Identification is based on the morphological characteristics of the organism. Although microscopic diagnosis of amebic infection is relatively quick and cheap, considerable expertise is required by the microscopist to differentiate the different amoeba. However the sensitivity and specificity of microscopy is poor (<60%) and can be confounded by false positive results due to misidentification.13,53 Also the trophozoites of E. ­histolytica degenerate rapidly in unfixed fecal samples and as such specimens should be preserved with a fixative to prevent morphological degradation.54 Due to the intermittent shedding of the parasite in fecal samples, for optimal microscopic detection of the organism, a minimum of three stool samples taken over no more than 10 days is recommended. Microscopy performed on aspirated material from ALA, are often negative for trophozoites, thus limiting the usefulness of microscopy as a diagnostic test for patients with ALA. The existence of E. dispar and E. moshkovskii in humans however, has made the definitive diagnosis of E. histolytica by microscopy impossible as the three species cannot be differentiated microscopically.13 48.1.5.2  Culture Various culture techniques have been successfully used for the diagnosis of amebiasis. Culture media include xenic (diphassic and monophasic) and axenic systems. Xenic cultures are defined as the growth of parasite in the present of an unidentified flora.55 Axenic cultures involve the cultivation of the parasites in the absence of any other metabolizing cells.55 In 1925 Boeck and Drbohlav were the first to successfully culture E. histolytica from clinical samples, using a xenic diphasic egg slope slant medium, with modifications of this media still used today.55 Other xenic culture systems developed for the cultivation of E. histolytica include; Balamuth medium,56 Jones’s medium,57 TYSGM-9,58 Locke-egg,55 and Robinson’s medium.59 Axenic cultivation of E. histolytica was first described by Diamond in 1961.60 Modifications to this medium were later developed and used in different research laboratories. Currently TYI-S-33 and YS-S medium are the most commonly used axenic media for the cultivation of E. histolytica.55 Various clinical specimens can be used for the culturing of E. histolytica including fecal samples, biopsy specimens, and liver abscess aspirates. The success rate for the cultivation of E. histolytica from clinical samples is approximately 50%.55 Culture is usually only performed in research laboratories

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due to the fact that the procedures are technically difficult, labor intensive and often unrewarding. Overgrowth of the medium is a major problem and cultures a have a significant false-negative rate. As such culture is not recommended for use in routine diagnostic laboratories. 48.1.5.3  Isoenzyme Analysis Isoenzyme analysis can differentiate Entamoeba species on the basis of zymodeme patterns.8 A zymodeme is defined as a group of ameba strains that share the same electrophoretic pattern and mobility for several enzymes (malic enzyme, hexokinase, glucose phostphate isomerase, phosphglucomutase isoenzyme).61 Isoenzyme analysis can differentiate E. histolytica from E. dispar and was considered the gold standard for diagnosing amebic infection prior to the development of DNA-based techniques. The technique does have a number of disadvantages. The test is difficult to perform, complex, costly, is time consuming and relies on the establishment of amoeba in culture. The process is not always successful, and cultivation of the amoeba may lead to selection bias. Zymodeme analysis is now restricted mainly to research laboratories and in the diagnostic lab it has been superseded by PCR-based methods. 48.1.5.4 Antibody Detection A number of different serological assays are commercially available for the detection of E. histolytica antibodies. Indirect hemagglutination, latex agglutination, complement fixation, indirect immunofluorescence assay, and enzymelinked immunosorbent assays have all been developed.13 Serological testing may be helpful from a diagnostic perspective in certain situations. In patients suspected of having extraintestinal disease, serological tests are warranted, however testing in patients with intestinal disease is normally not recommended. The serological tests are specific but have varying sensitivity depending on the presence or absence of invasive disease and the type of invasive disease. The sensitivity for serology is about 100% in patients with ALA62 and 82% with invasive intestinal disease.13 Antibody testing to diagnose carriage or non-invasive disease is unhelpful, as the sensitivity is low at around 8%.20 Serum IgG antibodies persist for years and antibody titers can remain high for years after successful therapy and/or eradication of the organism. This limits the usefulness of the test in areas where infection is endemic and people have been exposed to E. histolytica, due to the inability of serological tests to distinguish past from current infection.13 However, serological tests may be helpful from a diagnostic perspective in industrialized countries, where infections and exposure to E. histolytica is uncommon. 48.1.5.5 Antigen Detection Stool antigen assays are reported to outperform microscopy and to be as sensitive (80–85%) and specific (99%) as culture with isoenzyme analysis for the detection of E. histolytica.63 They are also relatively cheap, and easy

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to perform. Several kits are commercially available for the detection of E. histolytica antigens in fecal samples. These tests can either determine the presence of the E. histolytica/E. dispar group as being distinct from the rest of the Entamoeba species or they can actually differentiate E. histolytica from E. dispar. Most of these tests are in EIA formats. While most have reported excellent sensitivity and specificity there are a ­number of conflicting reports with researchers finding lower than expected sensitivities/specificities of the assays.13 The triage parasite panel (TTP) is an immunochromatographic assay for the simultaneous detection of E. histolytica/E. dispar, Giardia lamblia and Cryptosporidium parvum. Garcia et al. reported that the sensitivity and specificity of the TTP were 96 and 99.1%, respectively.64 However other studies have shown that the sensitivity of the assay was low (68.3%) when compared to the ProSpecT EIA test65 and when compared to PCR the TTP performed poorly.66 Antigen based ELISA kits that are specific for E. ­histolytica use monoclonal antibodies against the Gal/GalNAc-specific lectin (E. histolytica II, TechLab; Entamoeba CELISA, Cellabs,; ProspecT EIA, Remel Inc) or monoclonal antibodies against the serine-rich antigen of E. histolytica (Optimin S kit, Merlin Diagnostika). The TechLab antigen kit has been used over several years in different laboratories for the detection of E. histolytica in both endemic and nonendemic regions of the world. Results obtained using the TechLab antigen kit are conflicting with studies conducted in E. histolytica high endemic countries reporting high sensitivities between 95 and 100%.67,68 However, a recent study conducted in a high E. histolytica endemic region of northern Ecuador found that the Techlab E. histolytica II test performed poorly with a reported sensitivity of 14.3% and a specificity of 98.4% when compared to isoenzyme analysis.69 Stark et al. found the CELISA PATH kit showed 28% sensitivity and 100% specificity when PCR was used as a reference standard. In the same study the TechLab ELISA kit failed to identify any of the E. histolytica samples, which were positive by PCR.70 In low endemic settings the TechLab ELISA was documented to have a lower sensitivity than that of microscopy.71 Similar results were obtained when the TechLab ELISA was compared to real time PCR as a reference test.72 The ProspecT EIA (Remel Inc.) is a microplate EIA that while detects E. histolytica and E. dispar has the inability to distinguish the two. When compared to microscopy the sensitivity of this assay was reported between 73.5 and 78% while the specificity was 97.7 and 99%.73,74 When the assay was compared to culture and isoenzyme analysis the sensitivity and specificity were lower at 54.5 and 94%, respectively.69 Quantitative assessments have shown that EIA kits are 100–1,000 times less sensitive than PCR for detecting E. histolytica.70,75 These studies however used cell lysates of E. histolytica to calculate the analytical sensitivity of the ELISA’s, and it is not clear if the ELISA’s have comparable limits of detection of trophozoites, cysts, and cell lysates.

Molecular Detection of Foodborne Pathogens

Another disadvantage of antigen detection tests is that the antigens detected are denatured by fixation of the stool specimen, limiting the detection to fresh or frozen specimens.13 48.1.5.6  Molecular Diagnosis The PCR method for detection of E. histolytica is a powerful tool and offers excellent sensitivity and specificity. DNA-based tests for the diagnosis of amebiasis have already become the gold standard by which other diagnostic techniques (microscopy, antibody, and antigen detection) are measured. It should be noted however that PCR is susceptible to cross-contamination and false-negative results due to inhibitors of DNA polymerase in stool samples. Conventional PCR. To date a large number of conventional PCR methods have been described for the detection of E. histolytica targeting many different genes (Table 48.1). The initial studies used PCR to amplify the gene which encodes the 125-kDa surface antigen76 and this was further adapted by others to distinguish amongst the different Entamoeba species using restriction digestion.77 These initial experiments were performed on laboratory maintained control isolates of Entamoeba species. Primers targeting highly repetitive sequences present in E. histolytica were developed for PCR and used on DNA extracted from cultured trophozoites from feces; these primers amplify E. histolytica and E. dispar.78,79 The most widely assays for E. histolytica PCR target the 18S rDNA. These targets are present on multicopy, extrachromosomal plasmids in the ameba, making the 18S rDNA more easily detected and thus more sensitive than single copy gene targets.80 The consistent genetic diversity found in the 18S rDNA allows for the differentiation between E. histolytica and the closely related E. dispar and E. moshkovskii. PCR methods targeting this loci have proved to be highly sensitive and specific for detecting Entamoeba DNA.14,15,18,81–85 The 18S rDNA targets can be successfully amplified from either DNA extracted from laboratory cultured trophozoites or DNA extracted directly from feces. Both single round and nested PCR have been developed using 18S rDNA. A single round multiplex PCR assay was developed by Hamzah et al. for the detection and differential diagnosis of E. histolytica, E. dispar, and E. moshkovskii in humans.86 The PCR targeted the 18S rDNA by utilizing a conserved forward primer and reverse primers that are species specific to each of the three Entamoeba species. The assay was able to detect as little as 10 pg of DNA while exhibiting no cross-reactivity with other bacterial or protozoan parasites. Other targets include primers that amplify other regions of the extrachromosomal circular DNA.87,88 The 29-kDa/30kDa antigen gene was also used to distinguish between E. histolytica and E. dispar.89 This method was used with both DNA extracted from culture90 and direct fecal samples fixed in formalin.91,92 Protein encoding genes which have demonstrated polymorphisms in the coding region have also been utilized as targets for PCR. The serine-rich protein (SREPH) gene and the chitinase gene have both been used.93,94 In addition a novel PCR assay based on the hemolysin gene HLY6 was developed for the detection of E. histolytica DNA from

AAGCATTGTTTCTAGATCTGAG AAGAGGTCTAACCGAAATTAG

E-1 E-2

EH-1 EH-2

E-1 E-2 Eh-1 Eh-2

TAAGATGCACGAGAGCGAAA GTACAAAGGGCAGGGACGTA

EhP1 EhP2

Conventional PCR Tandem repeats in extrachromosomal circular rDNA Nested PCR SSU rRNA

Nested PCR SSU rRNA

CGATTTTCCCAGTTAGAAATTA CAAAATGGTCGTCGTCTAGGC

Eh5 Eh3

Conventional PCR SSU rRNA

TTTGTATTAGTACAAA GTA[A/G]TATTGATATACT AATGGCCAATTCATTCAATG TTTAGAAACAATGCTTCTCT

GTAACTTACTTAACCGGTAAAACATG TCTCTTCGTAACAAAGATCTAGACTC

ATG CAC GAG AGC GAA AGC AT GAT CTA GAA ACA ATG CTT CTC T

EntaF EhR

Conventional PCR SSU rRNA

GGCCAATTCATTCAATGAATTGAG CTCAGATCTAGAAACAATGCTTCTC

Sequence (5’–3’)

PSP5 PSP3

Primer/ Probe

Conventional PCR SSU rRNA

Assay Gene Target

Table 48.1 Conventional PCR Assays for E. histolytica Detection

900 bp

439 bp

132 bp

880 bp

166 bp

877 bp

Amplicon Size

First round: 96°C for 2 min, 30 cycles of 92°C for 60 1 min, 56°C for 1 min, 72°C for 90 sec, with a final extension at 72°C for 7 min Second round: 96°C for 2 min, 30 cycles of 92°C for 1 min, 48°C for 1 min, 72°C for 90 sec, with a final extension at 72°C for 7 min First round: 96°C for 2 min, 50 cycles of 92°C for 1 min, 43°C for 1 min, and 72°C for 90 sec Second round: 96°C for 2 min, 50 cycles of 92°C for 1 min, 62°C for 1 min, and 72°C for 90 sec

95°C for 5 min, 45 cycles of 95°C for 15 sec, 65°C for 15 sec, 72°C for 1 min, with a final extension at 72°C for 10 min 94°C for 3 min, 30 cycles of 90°C for 1 min, 58°C for 1 min, and 72°C for 1 min, with a final extension at 72°C for 7 min 94°C for 3 min, 40 cycles of 90°C for 1 min, 60°C for 1 min, 72°C for 2 min, with a final extension at 72°C for 10 min 30 cycles of 92°C for 1 min, 55°C for 1 min, and 70°C for 90 sec

Cycling Profile

0.4 μM

0.3 μM

0.4 µM

18 pmol

0.1 µM

0.4 µM

Primer Concentration

Detection Method

1.3% agarose gel stained with ethidium bromide

1.8% agarose gel stained with ethidium bromide

10% acrylamide gel stained with ethidium bromide

1% agarose gel stained with ethidium bromide

1.5% agarose gel stained with ethidium bromide

2% agarose gel stained with ethidium bromide

121

97

96

81

86

10

Reference

Entamoeba 683

684

both fecal and ALA samples and was shown to have excellent sensitivity and specificity.95 Nested PCR has also been developed in an attempt to increase the sensitivity of detection of E. histolytica. When utilizing a nested multiplex PCR for the simultaneous detection and differentiation of E. histolytica and E. dispar the sensitivity and specificity were 94% and 100%, respectively.96 A more recent study developed a nested multiplex PCR targeting the 18S rDNA for the differential detection of E. histolytica, E. dispar, and E. moshkovskii simultaneously in stool samples.97 This PCR demonstrated a sensitivity of 94% and a specificity of 100% for the demonstration of E. histolytica, E. dispar, and E. moshkovskii DNA in stool samples. ALA. An important and serious complication of intestinal infection with E. histolytica is the involvement of the liver (hepatic amebiasis).34 Hepatic amebiasis is usually diagnosed by the clinical picture (pain in the right upper quadrant and fever), ultrasound examination, and positive serology. However, none of these tests are definitive and the picture overlaps with pyogenic liver abscess caused by bacteria. Direct demonstration of E. histolytica by conventional microscopy and in vitro culture in pus obtained from ALA are often unsuccessful.13 PCR has been shown to be more reliable and a better alternative diagnostic test for ALA. A comparative study was done to verify the sensitivity of ten pairs of primers specific for detecting E. histolytica in stools.98 Of the ten pairs of previously published primers tested, two pairs of primers (PI+P2 and P11+P12) both targeting the extrachromosomal circular DNA of E. histolytica were found to give 100% sensitivity. Based on these results, the authors recommended that PCR assay can be successfully used to confirm the diagnosis of ALA with the primers identified.98 Previously Tachibana et al. used a PCR that employed primers specific for the gene encoding the 30 kDa molecule of E. histolytica.99 Liver abscess fluids (19 samples), from 14 patients with a presumptive amebic liver abscess, were examined microscopically and by the PCR method. Only two of the 19 samples were positive microscopically, whereas all 19 samples tested positive by PCR.99 A more recent study using another set of primers found the sensitivity and specificity of PCR for detecting ALA were 83% and 100%, respectively. The overall sensitivity of PCR was higher when compared to serological tests.100 Real-time PCR. A number of distinct real-time PCR (RT-PCR) protocols have been published for the identification and detection of E. histolytica (Table 48.2). RT-PCR offers several advantages over conventional PCR including elimination of post-PCR analysis reducing the risk of amplicon contamination, and faster cycling times resulting in reduced assay turn around times. RT-PCR also allows for the quantitation of organisms. These benefits do come at a cost with RT-PCR significantly more expensive than conventional PCR. Like conventional PCR assays the majority of the RT-PCR assays have targeted the rDNA. A SYBR green assay that did not utilize any specific probes, was described

Molecular Detection of Foodborne Pathogens

by Qvarnstrom et al.101 Blessmann et al. reported high sensitivity and specificity utilizing an RT-PCR assay performed with the LightCycler system using fluoresceinlabeled detection probes and primers amplifying a 310-bp fragment from the rDNA.102 The assay was able to detect as little as 0.1 parasite per gram of feces. Two RT-PCR assays were developed using Taqman probes,103 one targeting the SSU rDNA104 and another targeting the episomal repeats.105 An RT-PCR assay using a molecular-beacon targeting the 18S rDNA for use on fecal and ALA specimen has been described.106 An evaluation study conducted by Qvarnstrom et al., compared three E. histolytica real-time PCR techniques (Light cycler assay argetting the 18S rDNA, a Taqman assay targeting episomal repeats and a Taqman assay targeting the 18S rDNA) and a SYBR green RT-PCR method.101 The limits of detection and efficiency of each real-time PCR assay were determined and major differences in detection limits and assay performance were observed among the evaluated tests. Two of the assays could not reliably distinguish E. histolytica from E. dispar: the LightCycler assay and the TaqMan assay targeting episomal repeats. The LightCycler assay was found to be prone to false positives and clearly illustrates a lack of specificity of the primers. The TaqMan assay targeting episomal repeats also produced false results. Other publications have reported peculiar results concerning detection of E. dispar in conventional PCRs targeting these episomal-repeat sequences questioning the usefulness of this target for molecular testing.83,107 In conclusion, this study identified the TaqMan targeting the 18S rRNA gene as a superior real-time PCR assay for specific and quantitative diagnosis of amebiasis. The SYBR Green assay also performed well and offered a good alternative to the TaqMan assay.101

48.2  Methods 48.2.1 Reagents and Equipment Thermocycler Real-time PCR system DNA template that contains the region of the DNA fragment to be amplified One or more primers, which are complementary to the DNA regions at the 5′ and 3′ ends of the DNA region that is to be amplified DNA polymerase (e.g., Taq polymerase or another DNA polymerase with a temperature optimum at around 70°C), used to synthesize a DNA copy of the region to be amplified Deoxynucleotide triphosphates (dNTPs), from which the DNA polymerase builds the new DNA Buffer solution, which provides a suitable chemical environment for optimum activity and stability of the DNA polymerase Divalent cation, magnesium or manganese ions; generally Mg2+ is used, but Mn2+ can be utilized for

Molecular beacon

MGB-TaqMan probe SSU rRNA

TaqMan 2 Episomal repeats

Ehf Ehr Molecular beacon probe

Eh-S26C Eh-Ed-AS25 Eh-Ed-24-R Eh-Ed-25-D Ehd-239F Ehd-88R histolytica-96T histolytica-50F histolytica-132R histolytica-78T Ehd-239F Ehd-88R Histolytica-96T

LightCycler SSU rRNA

TaqMan 1 SSU rRNA

PSP5 PSP3

Primer/ Probe

SYBR Green realtime PCR SSU rRNA

Assay Gene Target

AACAGTAATAGTTTCTTTGGTTAGTAAAA CTTAGAATGTCATTTCTCAATTCAT Texas Red-GCGAGCATTAGTACAAAATGGCCAATTCATTCAGCTCGC-dR Elle

GTACAAAATGGCCAATTCATTCAACG GAATTGATTTTACTCAACTCTAGAG LC640-TCGAACCCCAATTCCTCGTTATCCp GCCATCTGTAAAGCTCCCTCTCCGA-FAM ATTGTCGTGGCATCCTAACTCA GCGGACGGCTCATTATAACA FAM-UCAUUGAAUGAAUUGGCCAUUU-BHQ1 CATTAAAAATGGTGAGGTTCTTAGGAA TGGTCGTCGTCTAGGCAAAATATT FAM-TTGACCAATTTACACCGTTGATTTTCGGA-BHQ1 CATTAAAAATGGTGAGGTTCTTAGGAA TGGTCGTCGTCTAGGCAAAATATT FAM-TTGACCAATTTACACCGTTGATTTTCGGA-BHQ1

GGCCAATTCATTCAATGAATTGAG CTCAGATCTAGAAACAATGCTTCTC

Sequence (5’–3’)

Table 48.2 Real-Time PCR Assays for E. histolytica Detection

134 bp

172 bp

83 bp

231 bp

307 bp

877 bp

Amplicon Size

45 cycles of 95°C for 15 sec, 55°C for 30 sec, and 72°C for 15 sec. The ramping of the machine was 3.3°C/sec in every step

95°C for 3 min, followed by 40 cycles of 95°C for 15 sec, 60°C for 30 sec, 72°C for 30 sec 95°C for 3 min, followed by 40 cycles of 95°C for 15 sec, 60°C for 30 sec, 72°C for 30 sec 95°C for 15 min followed by 40 cycles of 95°C for 15 sec, 60°C for 30 sec, and 72°C for 30 sec

95°C for 15 min, followed by 50 cycles of 95°C for 10 sec, 58°C for 10 sec, 72°C for 20 sec

95°C for 15 min, followed by 50 cycles of 95°C for 15 sec, 60°C for 1 min, 72°C for 1.5 min, 80°C for 30 sec + melting curve

Cycling Profile

0.5 µM of each primer/0.1 µM of probe 0.5 µM of each primer/0.1 µM of probe 6.25 pmol of each primer/1.75 pmol of probe 25 pmol of each primers/ 6.25 pmol of probe

0.5 µM of each primer/0.1 µM of each probe

0.1 µM of each primer

Primer/Probe Concentration

Detection Method

Fluorescence at the end of each extension plateau

Fluorescence at the end of each extension plateau Fluorescence at the end of each extension plateau Fluorescence at the end of each extension plateau

Fluorescence at the end of the 80°C incubation plateau and during the melting curve Fluorescence at the end of each annealing plateau

106

104

12

105

102

101

Reference

Entamoeba 685

686

PCR-mediated DNA mutagenesis, as higher Mn2+ concentration increases the error rate during DNA synthesis Monovalent cation potassium ions Real-time PCR probes PCR grade sterile H2O Pipettes Aerosol barrier filter tips

48.2.2 Sample Collection and Preparation Fecal samples are considered the most complex specimens for direct PCR testing because of the presence of PCR inhibitors such as heme, bilirubins, bile salts, and complex carbohydrates which are often co-extracted along with the target DNA.13 Therefore, optimization of the fecal DNA extraction procedure is critical. The transport and preservation conditions, of fecal samples, needs to be taken into consideration. A recent study showed that transportation of fecal samples containing E. histolytica at ambient temperature may result in the degeneration of parasite DNA, especially for the labile trophozoite stage.108 Therefore, the sensitivity of DNA assays using unpreserved samples is time-dependent. Specimens may be preserved by refrigeration or freezing. Many laboratories commonly use fixatives to preserve parasite morphology and for safe handling of specimens. Samples preserved in fixative may decrease the sensitivity of PCR with time.81 Several researchers have shown that formalin fixed samples can be used for DNA extraction.109 No reduction in the ability to perform PCR amplifications of E. histolytica DNA fixed in 1–10% formalin was noted for 7 days.110 However other researchers have shown formalin to interfere with the amplification.81 Consequently, if testing of fresh sample is not an option the freezing of a fresh specimen at –20ºC before extraction of DNA may be the best strategy as this does not affect the sensitivity of the molecular assays and allows for storage that is not time dependent.15 Recent, simple and effective methods for the isolation of DNA from feces have been developed. Commercially available DNA extraction kits are recommended due to the ease and success as they minimize extraction time and the DNA can be extracted directly from the feces without the need to culture the parasite. The following kits were used for the direct extraction of DNA from fecal samples: XTRAX DNA extraction kit111 (Gull laboratories, Salt Lake City, UT), Extract Master Fecal DNA extraction kit 9Epicenter Biotechnologies, WI) and the Genomic DNA Prep Plus kit.112 Of the methods for DNA extraction Qiagen kits have been used the most widely with the QIAamp stool kit now predominating. QIAamp tissue kit spin columns (Qiagen) have also been used for the extraction and purification of DNA from fecal samples.83 Various modifications including optimizing the duration and temperature of proteinase K digestion, adding additional wash steps, and treatment of the sample with 2% polyvinylpolypyrolidone have all been used.104,106 The QIAamp stool kit has proven to be the most widely

Molecular Detection of Foodborne Pathogens

accepted method for DNA extraction providing successful and reliable recovery of DNA from fecal material.15,16,18,70 Aspirated pus from ALA can also be tested by PCR and has shown excellent sensitivity and specificity.98,99 The following method is adapted from Verweij et al.104,113 for parasite DNA isolation from feces using QIAamp spin columns (QIAamp DNA minikit cat# 51306). (1) Suspend 0.1 g of feces in 200 µl of PBS with 2% polyvinylpolypyrolidone (PVPP**) in a microfuge tube. (2) Heat feces-suspension for 10 min. (3) Add 180 µl of tissue lysis buffer (ATL) and 20 µl of proteinase K from the QIAamp kit to the sample and mix by vortexing. (4) Incubate for 2 h (or overnight) at 55ºC in a heating block, after 1 h vortex briefly. (5) Add 400 µl of AL buffer to 400 µl of sample, mix thoroughly by vortexing. (6) Incubate for 70ºC for 10 min. (7) Centrifuge for 10 sec full speed and transfer the supernatant to a microfuge tube containing 400 µl of ethanol (96–100%), and mix thoroughly by vortexing. (8) Place a spin column in a 2-ml collection tube. Add 600 µl of the mixture from step 7 to the spin column without moistening the rim, close the cap and centrifuge at 10,000 rpm for 1 min. (9) Place spin column in a clean 2 ml collection tube a repeat with the rest of the mixture from step 7. (10) Place the spin column in a clean 2-ml collection tube. (11) Add 500 µl of AW1 buffer (from kit) to the spin column and centrifuge at 10,000 rpm for 1 min. (12) Empty the collection tube and add 500 µl of AW2 (from kit) buffer and centrifuge for 3 min at full speed. (13) Place the spin column in a clean 1.5 ml microfuge tube and elute DNA by adding 200 µl of AE buffer (from kit) and incubate for 1 min at room temperature. (14) Centrifuge at 8,000 rpm for 1 min. The eluted DNA is now ready for use in PCR.

48.2.3  Detection Procedure Principle: We present here a PCR assay that was developed by Troll et al. for specific detection of pathogenic E. histolytica.81 This assay employs forward primer Eh5 (5′-GTAACTTA CTTAACCGGTAAAACATG-3′) and reverse primer Eh3 (5′-TCTCTTCGTAACAAAGATCTAGACTC-3′)  derived from the small subunit (18S) rRNA gene for amplification of an 880 bp fragment from E. histolytica only. *

PVPP (sigma P6755) does not dissolve, and it stays in a suspension so regular stirring is needed when in use.

687

Entamoeba

Procedure: (1) Prepare PCR mixture (50 µl) containing 1× PCR buffer, 3 mM MgCl2, 200 µM dNTPs, 18 pmol of each primer (Eh5 and Eh3), 1.0 µl purified DNA, and distilled water (up to 50 µl). Include a tube containing the PCR mixture but not template DNA as negative control. (2) Conduct PCR amplification in a thermocycler with the following cycling program: one cycle of 94°C for 1 min; 40 cycles of 94°C for 1 min, 60°C for 1 min, and 72°C for 2 min; one cycle of 72°C for 10 min. (3) Separate the PCR product on 1% agarose with 0.5 µg/ml ethidium bromide. Visualize and photograph the product. Note: This method has been shown recently to be useful for identification of E. histolytica from stool samples.15

48.3 Conclusions and Further Perspectives It is now a little over 10 years since an expert committee on amebiasis suggested that there was a need to improve and advance the methods for diagnosis of E. histolytica. They acknowledged the difficulty of differentiating among this species and E. dispar; and this has been a major driving force behind research efforts, made more complex by the recent recognition of E. moshkovskii in human stool. It is therefore a rewarding experience to reflect on the achievements of the past ten years, and to recognize the advances that ELISA and PCR have contributed to this discipline. For the diagnostic laboratory, any diagnostic tests used for the diagnosis of amebiasis, needs to meet specific criteria in addition to the age old issue of cost. Determination of specificity and sensitivity of the test are paramount and the usefulness of the test within different settings raises considerations that may not be at first obvious. In developed countries such as Australia where infections with E. histolytica are not common, inclusion of gut parasites is often not the first choice in a differential diagnosis. Nevertheless increased awareness of amebiasis within the medical profession and the availability and provision of well characterized diagnostic tests by hospital laboratories will prevent amebiasis becoming a major public health problem114. Microscopy, despite all is limitations and especially the need for experienced, well trained microscopists, remains a relatively sensitive way to detect Entamoeba species in human stool. The need for provision of confirmatory tests, based on either ELISA or PCR, reduces the cost effectiveness of providing microscopy alone as a diagnosis. Fecal antigen detection tests represent an attractive follow up test because they are relatively quick to perform, easy to use and because they are often configured to the EIA format, they are not considered “unusual” to implement into the laboratory environment. In contrast technology such as real-time PCR, whilst probably the most effective diagnostic tool available at this moment in

tie, is still surrounded by a little “voodoo” and because of the cost involved (experienced staff, reagents, machines) is still unlikely to become the main stay for diagnosis. It is worth continuing the discussion on how to integrate technology into the diagnostic laboratory. Universities and colleges of science and technology have a roll to play in this through their educational programs and courses on professional development. Although molecular biology has been well integrated into many, if not most, educational courses in the biomedical sciences area, and graduates emerging from these courses are familiar with the terminology and jargon associated with molecular techniques, there is still resistance (cost) to introduce molecular technologies as routine diagnostic tests. Companies selling reagents and equipment must shoulder some of the blame here, since the prohibitive cost associated with technology such as real-time PCR, means the services are simply not provided. In this modern world where diagnosis can aid in the prevention of life threatening diseases, the failure to provide diagnostic services is almost unethical. How do we resolve this situation? As always technology may provide a solution to this dilemma.115 The detection of many pathogens in a single assay is an achievable goal in this day and age, and some simple tests are already commercially available. One can imagine, however, that chip-based technology may ultimately provide a means to detect all pathogens currently assayed for by diagnostic labs in addition to Entamoeba species.116,117 The added attraction in this approach is that it is also possible to include tests that monitor pathogenesis and disease occurrence at the same time. Once again arrays for numerous cellular pathways are commonly used by research laboratories, and their incorporation into tests for pathogens, should provide a powerful approach for diagnosis of pathogen and disease at the same time. What role is there for the immunoassay in a world based on molecular diagnosis? Nanotechnology may also provide a future for the immunoassay and the development of biosensors as immunosensors may eventually come forth with useful tools.118 Indeed advances in the detection of bacteria using immunosensors have recently been reviewed119 and the data suggests this technology may yet evolve into an efficient high throughput device for use in diagnostic labs.120 Ironically parasites that infect humans do not yet appear to be included in immunosensor prototypes at this moment in time.121 Finally, whatever the future holds for diagnosis of parasitic infections such as those caused by E. histolytica, it must be remembered that such tests require applicability in developing countries, especially those where amebiasis is endemic. This raises further challenges, not only of cost, but also how diagnostic approaches can be integrated into management of public health. Such issues, unlike advances in technology, are frequently difficult for governments to manage at a local level.

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3. Anonymous. WHO News and activities, Entamoeba taxonomy. Bull. WHO, 75, 291, 1997. 4. Losch, F.A. Massive development of amebas in the large intestine. Am. J. Trop Med. Hyg., 24, 383, 1875. 5. Councilman, W.T., and LaFleur, H.A. Amebic dysentery. Johns Hopkins Hosp. Rep., 2, 395, 1891. 6. Brukner, D.A. Amebiasis. Clin. Microbiol. Rev., 5, 356, 1992. 7. Walker, E.L., and Sellards, A.W. Experimental entamoebic dysentery. Philipp. J. Sci., 8, 253. 1913. 8. Sargeaunt, P.G., Williams J.E. and Grene, J.D. The differentiation of invasive and non-invasive Entamoeba histolytica by isoenzyme electrophoresis. Trans. R. Soc. Trop. Med. Hyg., 72, 519, 1978. 9. Clark, C.G. and Diamond, L.S., Entamoeba histolytica: an explanation for the reported conversion of “nonpathogenic” amebae to the pathogenic form, Exp. Parasitol., 77, 456, 1993. 10. Diamond, L.S. and Clark, C.G. A redescription of Entamoeba histolytica Schaudin, 1903 (emended Walker, 1911) separating it from Entamoeba dispar Brumpt, 1925. J. Eukaryotic. Microbiol., 40, 340, 1993. 11. Tanyuksel, M. and Petri, W.A., Jr. Laboratory diagnosis of Amebiasis. Clin. Microbiol. Rev., 16, 713, 2003. 12. Verweij, J.J. et al. Entamoeba histolytica infections in captive primates. Parasitol. Res., 90, 100, 2003. 13. Fotedar, R. et al. Laboratory diagnostic techniques for Entamoeba species. Clin. Microbiol. Rev., 20, 511, 2007. 14. Clark, C.G. and Diamond, L.S. Intraspecific variation and phylogenetic relationships in the genus Entamoeba as revealed by riboprinting. J. Eukaryot. Microbiol., 44, 142, 1997. 15. Fotedar, R. et al. Detection and identification of Entamoeba histolytica, Entamoeba dispar and Entamoeba moshkovskii in stool samples from an Australian population. J. Clin. Microbiol., 45, 1035, 2007. 16. Fotedar, R. et al. Entamoeba moshkovskii infections in Sydney, Australia. Eur. J. Clin. Microbiol. Infect. Dis., 27, 133, 2008. 17. Ali, I.K. et al. Entamoeba moshkovskii infections in children, Bangladesh. Emerg. Infect. Dis., 9, 580, 2003. 18. Stark, D. et al. Prevalence of enteric protozoa in HIV-positive and HIV-negative men who have sex with men from Sydney, Australia. Am. J. Trop. Med. Hyg., 76, 549, 2007. 19. Dobell, C. Research on the intestinal protozoa of monkeys and man. Parasitology, 20, 357, 1928. 20. World Health Organization. World Health Organization/Pan American Health Organization/UNESCO report of a consultation of experts on amebiasis. Wkly. Epidemiol. Rec., 72, 97, 1997. 21. Hutson, C.D. and Petri, W.A. Jr. Amebiasis: clinical implications of the recognition of Entamoeba dispar. Curr. Infect. Dis. Rep., 1, 441, 1999. 22. Walsh, J.A. Problems in recognition and diagnosis of amebiasis: estimation of the global magnitude of morbidity and mortality. Rev. Infect. Dis., 8, 228, 1986. 23. Petri, W.A. Jr. Recent advances in amebiasis. Crit. Rev. Clin. Lab. Sci., 33, 1, 1996. 24. Leroy, A.G. et al. Contact-dependent transfer of the galactosespecific lectin of Entamoeba histolytica to the lateral surface of enterocytes in culture. Infect. Immun., 63, 4253, 1995. 25. Stanley, S.L. et al. Role of the Entamoeba histolytica cysteine proteinase in amebic liver abscess formation in severe combined immunodeficient mice. Infect. Immun., 63, 1587, 1995.

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Entamoeba 53. Pillai, D.R. and Kain, K.C. Immunochromatographic stripbased detection of Entamoeba histolytica-E. dispar and Giardia lamblia coproantigen. J. Clin. Microbiol., 37, 3017, 1999. 54. Proctor, E.M. Laboratory diagnosis of amebiasis. Clin. Lab. Med., 11, 829, 1991. 55. Clark, C.G., and Diamond, L.S. Methods for cultivation of luminal parasitic protists of clinical importance. Clin. Microbiol. Rev., 15, 329, 2002. 56. Balamuth, W. Impoved egg yolk medium for the cultivation of Entamoeba histolytica and other intestinal protozoa. Am. J. Clin. Pathol., 16, 380,1946. 57. Jones, W.R. The experimental infection of rats with Entamoeba histolytica; with a method for evaluating the anti-amoebic properties of new compounds. Ann. Trop. Med. Parasitol., 40, 130,1946. 58. Diamond, L.S. A new liquid medium for xenic cultivation of Entamoeba histolytica and other lumen dwelling protozoa. J. Parasitol., 68, 958,1982. 59. Robinson, G.L. The laboratory diagnosis of human parasitic amoebae. Trans. R. Soc. Trop. Med. Hyg., 62, 285, 1968. 60. Diamond, L.S. Axenic cultivation of Entamoeba histolytica, Science, 134, 336, 1961. 61. Sargeaunt, P.G. et al. The reliability of Entamoeba histolytica zymodemes in clinical laboratory diagnosis. Arch. Investig. Med., 18, 69, 1987. 62. Zengzhu, G. et al. Analysis by enzyme-linked immunosorbent assay and PCR of human liver abscess aspirates from patients in China for Entamoeba histolytica. J. Clin. Microbiol., 37, 3034,1999. 63. Haque, R. et al. Rapid diagnosis of Entamoeba infection by using Entamoeba and Entamoeba histolytica stool antigen detection kits. J. Clin. Microbiol., 33, 2558,1995. 64. Garcia, L.S., Shimizu, R.Y. and Bernard C.N. Detection of Giardia lamblia, Entamoeba histolytical Entamoeba dispar, and Cryptosporidium parvum antigens in human fecal specimens using the Triage parasite panel enzyme immunoassay. J. Clin. Microbiol., 38, 3337, 2000. 65. Pillai, D.R. et al. Entamoeba histolytica and Entamoeba dispar: epidemiology and comparison of diagnostic methods in a setting of nonendemicity. Clin. Infect. Dis., 29, 1315, 1999. 66. Levia, B. et al. Overdiagnosis of Entamoeba histolytica and Entamoeba dispar in Nicaragua: a microscopic, triage parasite panel and PCR study. Arch. Med. Res., 37, 529, 2006. 67. Haque, R. et al. Entamoeba histolytica and Entamoeba dispar infection in children in Bangladesh. J. Infect. Dis., 175, 734, 1997. 68. Solaymani-Mohammadi, S. et al. Comparison of a stool ­antigen detection kit and PCR for diagnosis of Entamoeba histolytica and Entamoeba dispar infections in asymptomatic cyst passers in Iran. J. Clin. Microbiol., 44, 2258, 2006. 69. Gatti, S., et al. Amebic infections due to the Entamoeba ­histolytica-Entamoeba dispar complex: a study of the incidence in a remote rural area of Ecuador. Am. J. Trop. Med. Hyg., 67, 123, 2002. 70. Stark, D. et al. Comparison of stool antigen detection kits with PCR for the diagnosis of amebiasis. J. Clin. Microbiol., 46, 1678, 2008. 71. Gonin, P. and Trudel, L. Detection and differentiation of Entamoeba histolytica and Entamoeba dispar isolates in clinical samples by PCR and enzyme-linked immunosorbent assay. J. Clin. Microbiol., 41, 237, 2003.

689 72. Visser, L.G. et al. Diagnostic methods for differentiation of Entamoeba histolytica and Entamoeba dispar in carriers: performance and clinical implications in a non-endemic setting. Int. J. Med. Microbiol., 296, 397, 2006. 73. Jelinek, T. et al. Evaluation of an antigen-capture enzyme immunoassay for detection of Entamoeba histolytica in stool samples. Eur. J. Clin. Microbiol. Infect. Dis., 15, 752, 1996. 74. Ong, S.J. et al. Use of the ProSpecT microplate enzyme immunoassay for the detection of pathogenic and non-­pathogenic Entamoeba histolytica in fecal specimens. Trans. R. Soc. Trop. Med. Hyg., 90, 248, 1996. 75. Mirelman, D., Nuchamowitz, Y., and Stolarsky, T. Comparison of use of enzyme-linked immunosorbent assay-based kits and PCR amplification of rRNA genes for simultaneous detection of Entamoeba histolytica and E. dispar. J. Clin. Microbiol., 35, 2405, 1997. 76. Edman, U. et al. Characterisation of an immuno-dominant variable surface antigen from pathogenic and non-pathogenic Entamoeba histolytica. J. Exp. Med., 172, 879, 1990. 77. Tannich, E. and Burchard, G.D. Differentiation of pathogenic from non-pathogenic Entamoeba histolytica by restriction fragment analysis of a single gene amplified in vitro. J. Clin. Microbiol., 29, 250, 1991. 78. Garfinkel, L. et al. DNA probes specific for Entamoeba histolytica possessing pathogenic and nonpathogenic zymodemes. Infect. Immun., 57, 926, 1989. 79. Romero, J.L. et al. Use of polymerase chain reaction and nonradioactive DNA probes to diagnose Entamoeba histolytica infection. Arch. Med. Res., 23, 277, 1992. 80. Bhattacharya, S. et al., Circular DNA of Entamoeba histolytica encodes ribosomal RNA. J. Protozool., 36, 455, 1989. 81. Troll, H., Marti, H. and Weiss, N. Simple differential detection of Entamoeba histolytica and Entamoeba dispar in fresh stool specimens by sodium acetate-acetic acid-formalin concentration and PCR. J. Clin. Microbiol., 35, 1701, 1997. 82. Ramos, F. et al. Prevalence of Entamoeba histolytica and Entamoeba dispar in a highly endemic rural population. Arch. Med. Res., 31, 34, 2000. 83. Verweij, J. J. et al. Differentiation of Entamoeba histolytica and Entamoeba dispar cysts using polymerase chain reaction on DNA isolated from faeces with spin columns. Eur. J. Clin. Microbiol. Infect. Dis., 19, 358, 2000. 84. Heckendorn, F. et al. Species-specific field testing of Entamoeba spp. in an area of high endemicity. Trans. Roy. Soc. Trop. Med. Hyg., 96, 521, 2002. 85. Ramos, F. et al. Entamoeba histolytica and Entamoeba ­dispar: prevalence infection in a rural Mexican community. Exp. Parasitol., 110, 327, 2005. 86. Hamzah, Z. et al. Differential detection of Entamoeba histolytica, Entamoeba dispar, and Entamoeba moshkovskii by a single-round PCR assay. J. Clin. Microbiol., 44, 3196, 2006. 87. Acuna-Soto, R. et al. Application of the polymerase chain reaction to the epidemiology of pathogenic and nonpathogenic Entamoeba histolytica. Am. J. Trop. Med. Hyg., 48, 58, 1993. 88. Britten, D. et al. An improved colorimetric PCR-based method for the detection and differentiation of Entamoeba histolytica and Entamoeba dispar. J. Clin. Microiol., 35, 1108, 1997. 89. Tachibana, H. et al. Distinguishing pathogenic isolates of Entamoeba histolytica by polymerase chain reaction. J. Infect. Dis., 164, 825, 1991. 90. Tachibana , H. et al. Asymptomatic cyst passers of Entamoeba histolytica but not Entamoeba dispar in institutions for the mentally retarded in Japan. Parasitol. Int., 49, 31, 2000.

690 91. Rivera, W.L. et al. Differentiation of Entamoeba histolytica and E. dispar DNA from cysts present in stool specimens by polymerase chain reaction: its field application in the Philippines. Parasitol Res., 82, 585, 1996. 92. Rivera, W.L., Santos S.R. and Kanbara, H. Prevalence and genetic diversity of Entamoeba histolytica in an institution for the mentally retarded in the Philippines. Parasitol. Res, 98, 106, 2006. 93. Ayeh-Kumi, P.F. et al. Entamoeba histolytica: genetic diversity of clinical isolates from Bangladesh as demonstrated by polymorphisms in the serine-rich gene. Exp. Parasitol., 99, 80, 2001. 94. Freitas, M.A. et al. A single step duplex PCR to distinguish Entamoeba histoylytica from Entamoeba dispar. Parasitology, 128, 625, 2004. 95. Zindrou, S. Specific detection of Entamoeba histolytica DNA by hemolysin gene targeted PCR. Acta Trop., 78, 117, 2001. 96. Nunez, Y.O. et al. Multiplex polymerase chain reaction amplification and differentiation of Entamoeba histolytica and Entamoeba dispar DNA from stool samples. Am. J. Trop. Med. Hyg., 64, 293, 2001. 97. Khairnar, K. and Parija, S.C. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar DNA in stool samples. BMC Microbiol., 7, 47, 2007. 98. Zaman, S. et al. Direct amplification of Entamoeba histolytica DNA from amoebic liver abscess pus using polymerase chain reaction. Parasitol. Res., 86, 724, 2000. 99. Tachibana, H. et al. Detection of pathogenic Entamoeba histolytica DNA in liver abscess fluid by polymerase chain reaction. Int. J. Parasitol., 22, 1193, 1992. 100. Khan, U. et al. Detection of Entamoeba histolytica using polymerase chain reaction in pus samples from amebic liver abscess. Indian J. Gastroenterol., 25, 55, 2006. 101. Qvarnstrom, Y. et al. Comparison of real-time PCR protocols for differential laboratory diagnosis of amebiasis. J. Clin. Microbiol., 43, 5491, 2005. 102. Blessman, J. et al. Real-time PCR for detection and differentiation of Entamoeba histolytica and Entamoeba dispar in fecal samples. J. Clin. Microbiol., 40, 4413, 2002. 103. Kebede, A. et al. The use of real-time PCR to identify Entamoeba histolytica and E. dispar infections in prisoners and primary-school children in Ethiopia. Ann. Trop. Med. Parasitol., 98, 43, 2004. 104. Verweij, J.J. et al. Simultaneous detection of Entamoeba ­histolytica, Giardia lamblia, and Cryptosporidium parvum in faecal samples by using multiplex real-time PCR. J. CLin. Microbiol., 42, 1220, 2004. 105. Verweij, J.J. et al. Short communication: Prevalence of Entamoeba histolytica and Entamoeba dispar in northern Ghana. Trop. Med. Int. Health., 8, 1153, 2003.

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49 Encephalitozoon and Enterocytozoon Jaco J. Verweij

Leiden University Medical Center

Dongyou Liu

Mississippi State University

Contents 49.1 Introduction.................................................................................................................................................................... 691 49.1.1 Classification and Morphology......................................................................................................................... 691 49.1.2 Biology and Epidemiology................................................................................................................................ 692 49.1.3 Laboratory Diagnosis........................................................................................................................................ 693 49.1.3.1 Conventional Techniques................................................................................................................. 693 49.1.3.2 Molecular Techniques...................................................................................................................... 694 49.2 Methods.......................................................................................................................................................................... 695 49.2.1 Sample Preparation........................................................................................................................................... 695 49.2.2 Detection Procedures . ..................................................................................................................................... 696 49.2.2.1 Mutiplex Nested PCR20. .................................................................................................................. 696 49.2.2.2 Real-Time PCR25............................................................................................................................. 696 49.2.2.3 Genotyping of Enterocytozoon bieneusi on the ITS Region........................................................... 697 49.3 Conclusion and Future Perspectives .............................................................................................................................. 698 References.................................................................................................................................................................................. 698

49.1  Introduction 49.1.1 Classification and Morphology Encephalitozoon and Enterocytozoon are important members of a group of unicellular, small obligate intracellular, spore-forming organisms commonly known as “microsporidia,” which are classified under the phylum Microspora, subkingdom Protozoa, kingdom Protista. Being true eukaryotes, microsporidia possess a membrane-bound nucleus and an intracytoplasmic membrane system, and show chromosome separation on mitotic spindles. On the other hand, by possessing 70S ribosomes (with ribosomal subunits 30S and 50S containing prokaryote-sized 16S and 23S rRNAs without separate 5.8S rRNA, which is considered as an important eukaryotic feature), simple Golgi membranes, relatively small genome (about 2–7 megabase pairs), chitin and trehalose in the spore wall, and by being free of mitochondria or peroxisomes, microsporidia resemble fungi (especially the zygomycetes).1 Recent phylogenetic analyses of the smallsubunit (SSU) rRNA, the largest subunit of RNA polymerase II, TATA-box-binding protein, glutamyl tRNA synthase, αand β-tubulin and mitochondrial Hsp70 (a heat shock protein or chaperonin) gene sequences in microsporidia provide further evidence that microsporidia and fungi are close relatives, which may have similarly evolved degeneratively from higher forms.2,3 An extensive phylogenetic analysis using eight genes

placed microsporidia within the fungal clade, as a sister to a combined ascomycetes and basidiomycetes clade.4 Based on the presence or absence of an outer membrane surrounding the organisms, microsporidia can be separated into the Pansporoblastina (with membrane) and the Apansporoblastina (without membrane). According to the differences in chromosome cycles, microsporidia are divided into the Dihaplophasea (with a diplokaryon containing two adjoined nuclei in a single unit), and the Haplophasea (with unpaired nuclei).5 To date, over 140 genera and 1200 species that infect a wide variety of invertebrates and vertebrates have been identified in the phylum Microspora.6 Among these, at least 14 different species belonging to seven genera (i.e., Enterocytozoon, Encephalitozoon (syn. Septata), Pleistophora, Trachipleistophora, Vittaforma, Nosema, and Brachiola) along with some unclassified microsporidia have been associated with human ailments, particular in individuals with suppressed immune status.7 Among these, Encephalitozoon intestinalis, Enterocytozoon bieneusi, Nosema species, Pleistophora species, Vittaforma corneae (syn. Nosema corneum) have been isolated in water or fish, and pose risk as human waterborne, and foodborne pathogens.8 As microsporidia are obligate intracellular organisms that are unable to undertake active metabolic processes outside of the host cell, they are dependent on hosts for nutrients and 691

692

maintenance of their life cycle, which displays three ­distinct stages: spore, merogony, and sporogony. Usually measured 1.0–3.0 × 1.5–4.0 μm, microsporidial spores possess a thick wall composed of a proteinaceous outer layer called the exospore, a chitinous inner layer called the endospore, and a plasma membrane covering the cytoplasm. Within the cytoplasm (sporoplasm) are located the nucleus, a posterior vacuole, the polaroplast membranes, and the unique extrusion apparatus, which is composed of a coiled polar filament and a characteristic anchoring disc. Microsporidial nucleus may assume as a monokaryon (single nucleus, which is found in Encephalitozoon, Enterocytozoon, Pleistophora, and Trachipleistophora), or as a diplokaryon (having two adjoined nuclei in a single unit, which is found in Brachiola, Nosema, and Vittaforma). Upon ingestion or inhalation by a suitable host, which is often accompanied with a change in pH or osmotic pressure, the polar filament moves out of the anterior end of the spore, penetrates the host cell to inoculate the infective sporoplasm, which then grows to become a meront. Being round, irregular, or elongated simple cells without clearly differentiated cytoplasm, meronts undergo divisions by binary or multiple fissions called merogony. Multinucleated plasmodial forms may also appear as a result of nuclear division without cell division. In the next stage (i.e., sporogony), meronts turn into sporonts with a characteristic dense surface coat, which later forms the exospore layer of the spore wall. Also replicating by binary or multiple fissions, sporonts divide into ovoid-shaped sporoblasts that subsequently develop into mature spores (which are infective and relatively tolerant of environmental conditions) by synthesis of spore organelles.3

49.1.2  Biology and Epidemiology Microsporidial spores normally gain entry to susceptible host cells via active invasion and/or endocytosis. In the active invasion mode, microsporidial spores are activated upon exposure to a number of external stimuli, leading to the breakdown of sporoplasm compartmentalization and an increase in osmotic pressure inside the spore. This change results in the expansion of the posterior vacuole, and triggers eversion of the polar tube (which is made up of four to 30 coils around the sporoplasm in the spore) through the thinnest region of the spore. As the uncoiled polar tube measures between 50 μm and 500 μm (about 100 times of the normal length of the spore), it facilitates rapid and direct injection of the sporoplasm into the susceptible host cell. The process of spore activation and polar tube discharge (injection of the infective sporoplasm into the host cell) takes less than 2 sec. With the infective sporoplasm being released into the cytoplasm of the host cell and covered by a new plasma membrane derived from the polaroplast, the remaining empty spore and polar tube are degraded. In the cytoplasm, the sporoplasm matures into a meront, which goes through asexual reproduction (merogony) to amount to 50 to several hundred meronts in a single cell. The meronts next develop into sporonts (showing a typical dense surface coat), which further

Molecular Detection of Foodborne Pathogens

divide into ovoid-shaped sporoblasts and then become mature spores. Following the burst of the host cells, the infective spores are released. In the endocytosis mode, a microsporidial spore is passively taken up by the host cell (mediated by directed, actin-dependent phagocytosis) and first located in a phagosome, where the spore releases its infective sporoplasm into the host cell cytoplasm with the help of its polar tube and action of the host cell lysosomes. The sporoplasm then develops into meronts and sporonts as in the active invasion mode.9 A recent study indicates that in the endocytosis mode, the microsporidial spore attaches (or is engulfed by) to macrophages within the first 3–6 h, where the spore organizes into internal parasitophorous vacuoles at 24 h, replicates within the cytoplasm of macrophages at 72 h, and produces mature spores at 120 h post infection.10 Microsporidia are capable of infecting a wide range of invertebrates and vertebrates, from which microsporidial spores are discharged into the environment via the feces, urine or respiratory secretions. With a chitinous wall, microsporidial spores are relatively resistant to various environmental conditions. For example, drying (dehydration), incubation in distilled water or acid/alkali buffer (e.g., pH 4/pH 9), or freezing and thawing fail to kill all E. cuniculi, E. hellem, E. intestinalis spores.9 Thus, microsporidial spores remain infectious in the water, and other environments, and are readily transmitted to susceptible hosts via fecal-oral or oral-oral route, inhalation of aerosols, or ingestion of food contaminated with fecal material. Direct human-to-human transmission of intestinal microsporidiosis is also observed in some rare cases involving men who have sex with men.11 Enterocytozoon bieneusi and Encephalitozoon intestinalis represent the most common causes of microsporidial infections in humans. In immunologically competent human hosts such as travelers, microsporidia infections such as E. bieneusi and E. intestinalis may pass unnoticed or usually present with mild clinical signs which may include self-limiting diarrhea.7 Entering via the cornea, Vittaforma corneae, Brachiola algerae, and Nosema species tend to induce ocular infections, and contribute to blurred vision and corneal rupture in healthy individuals. In immune-compromised individuals, microsporidia such as E. bieneusi and E. intestinalis produce persistent diarrhea along with fever, nausea, fatigue, weight loss, wasting disease, and death. Additionally, Encephalitozoon, Trachipleistophora, and Pleistophora species can take advantage of host’s suppressed immune status and cause encephalitis, sinusitis, keratoconjunctivitis, hepatitis, myositis, nephritis, encephalitis, pneumonia, or peritonitis. Malnutrition in children may lead to persistent diarrhea upon E. bieneusi infection.6,7,8 Microsporidia infecting humans and animals appear to belong to genetically distinct strains. For example, three strains (I, II, and III) of E. cuniculi are known, which are also named “rabbit strain,” “mouse strain” (infecting both rodents and blue foxes), and “dog strain”.12 At the molecular level, E. cuniculi strain I possesses three copies of the repeated element 5′-GTTT-3′ in the internal transcribed spacer (ITS), strain II has two such repeats, and strain III harbors four.

Encephalitozoon and Enterocytozoon

In terms of geographical distribution, E. cuniculi strain I ­infecting rabbits is reported in three continents (America, Australia, and Europe), strain II is identified in Europe only, and strain III is found in dogs in America and South Africa as well as in tamarin colonies from zoos in Europe. E. cuniculi strain I (“rabbit strain”) has been shown to be responsible for microsporidiosis in immunosuppressed individuals in Europe and strain III (“dog strain”) in the Americas.6–8,12 Similarly, using the ITS sequence as genetic marker, three genotypes (1, 2, and 3) are recognized in E. hellem, with genotype 1 accounting for most of E. hellem isolates characterized so far from patients in the United States and in Europe; genotypes 2 and 3 and 2b consisting of a small number of human-derived E. hellem isolates from Switzerland and Tanzania. 6,8,13,14 There is clear evidence that some E. bieneusi genotypes are host-specific, whereas others are zoonotic because they have been found to infect both humans and animals. For example, through examination of a 243-bp nucleotide sequence of the ITS region of the rRNA gene, over 50 E. bieneusi genotypes have been identified to date in farm and companion animals including pigs, cattle, cats, dogs, and goats; in wildlife including beavers, muskrats, otters, foxes, and raccoons; and in laboratory rhesus macaques. On the other hand, of the 17 E. bieneusi genotypes that have been shown to infect humans, four are clearly zoonotic and capable of infecting other vertebrate hosts, while genotypes A, B, and C dominate clinical isolations from humans. Interestingly, analyses of the ITS sequences from human-infecting­infecting E. bieneusi suggest that E. bieneusi genotypes can be clustered according to host species. Indeed, examination of the ITS region led to the identification of 11 E. bieneusi genotypes (BEB1/J/PtEbX/CEbB, BEB2/I/PtEbXI/CEbE, BEB3, BEB4, BEB6, BEB7, N, M, CEbA, CEbD, and CEbF) that demonstrate specificity for cattle. However, several other E. bieneusi genotypes such as K (or IV/BEB5/Peru2), D (or PigEBTS9, Peru9, WL8, and CEbC) have been recovered from humans and animals.8,15

49.1.3 Laboratory Diagnosis As microsporidia are small organisms that tend to cause nonspecific early symptoms, diagnosis of microsporidiosis is dependent on the application of various phenotypic or genotypic laboratory procedures. Whereas conventional techniques (e.g., microscopy, in vitro culture, in vivo assays, and serological tests) have proven useful for the phenotypic characterization of microsporidia, molecular techniques have been established as the preferred methods for improved diagnosis of microsporidiosis in recent years. 49.1.3.1  Conventional Techniques Microscopy. Microsporidial spores in biopsy tissues, body fluids (urine, nasal discharge, sinus aspirates, sputum, bronchoalveolar lavage fluid, duodenal or bile juice, cerebrospinal fluid), or stool samples are often detected under light ­m icroscopy with the help of special stains such as chemofluorescent (optical white) agents, staining the chitin wall of

693

Figure 49.1  Enterocytozoon bieneusi spores in optical white staining. Courtesy of Eric A.T. Brienen, LUMC, Leiden.

the mature spores (Figure 49.1),16 or concentrated chromotrope stain (modified trichrome).17 Occasionally spores are observed with periodic acid-Schiff (PAS) stain, silver stains (e.g., Warthin-Starry), acid-fast stains, a chromotrope-based stain, or chemofluorescent agents. However, identification of microsporidia meront and sporont stages in tissue sections processed with Giemsa, hematoxylin, or eosin stain can be challenging. Given the presence of chitin in the endospore layer, microsporidial spores (showing violet to purple color) are differentiated from lysosomes, mucin droplets and other debris with a combination of Gram and modified Warthin–Starry stains, or tissue Gram stains (Brown–Brenn or Brown–Hopps). For spore detection in ultrathin plastic sections, it is advisable to use methylene blue-azure II-basic fuchsin or toluidine blue stain.17 Nonetheless, light microscopy does not permit accurate species determination, which is important for selection of appropriate chemotherapy. The unique ultrastructure of the characteristic coiled polar tube and other important structural features of the microsporidial spores can be visualized using electron microscopy (EM). Together with the knowledge of their proliferative forms, method of division, and nature of the host cell-parasite interface, microsporidial spores can be then determined to the genus or even species level. EM has long been regarded as the gold standard for confirmation of microsporidial identity. Nevertheless, identification of microsporidial spores using EM is time-consuming and expensive. Furthermore, it has become increasingly apparent that morphological examination is insufficient for precise determination of microsporidial species or subspecies without detailed assessment of their antigenic and molecular characteristics using other techniques. In vitro culture. Apart from E. bieneusi, which is one of the most common human microsporidial pathogens, and yet only survives in culture for a short-term, most microsporidia from human hosts have the capacity to continuously grow in various established cell lines. The most commonly used cell lines for microsporidial in vitro culture are those derived from monkey and rabbit kidney (Vero and RK-13),

694

human fetal lung fibroblast (MRC-5), and the Madin–Darby canine kidney (MDCK). In addition to providing a confirmatory diagnosis of microsporidial spores, in vitro culture techniques have proven crucial for the elucidation of microsporidial biology, host-parasite interplay, and pathogenesis; for the development and evaluation of serological diagnostic reagents; and for the assessment of the chemotherapy, disinfection, and sterilization regimens that are designed to treat and control microsporidiosis. Despite their time-consuming and laborious nature, in vitro culture techniques continue to play a supporting role in the laboratory confirmation of infections resulting from existing or new microsporidial species, along with microscopic examination, serological, and molecular analyses. Serological tests. A large number of serological methods have been developed for specific detection of microsporidiaspecific antibodies and antigens. These include carbon immunoassay, immunofluorescent antibody staining, complement fixation, enzyme-linked immunosorbent assay (ELISA), and western immunoblot, etc. The indirect immunofluorescence test and ELISA have been widely used for the diagnosis of microsporidiosis in immunological competent laboratory animals. In particular, monoclonal antibodies targeting the surface proteins and polar tubes of microsporidial spores have been employed for specific detection of E. hellem, E. cuniculi, or E. intestinalis. However, considering that microsporidiosis frequently occurs in individuals with compromised immune status, who tend to display poor immune response to the invading pathogens, the benefit of using serologic methods to diagnose microsporidiosis in this most susceptible population group is severely limited. In vivo assays. Although not suitable for patient diagnosis, animal models do offer a valuable approach in the study of microsporidial infections that is not easily obtainable with in vitro culture and other related techniques. For example, examination of route of transmission and host innate and acquired immune responses against microsporidia is impossible without conducting in vivo experiments. Similarly, development, and production of laboratory diagnostic reagents (e.g., monoclonal and polyclonal antibodies) are reliant on laboratory animals. Moreover, evaluation of therapeutic drugs and candidate vaccines against microsporidial infections is contingent on first obtaining some critical data concerning their toxicity and efficacy from animal models. Mice (e.g., BALB/c, athymic C57Bl/6, and SCID), rabbits and rhesus macaque monkeys represent some of the frequently applied animal models for the analysis of microsporidial infections such as E. cuniculi, E. hellem, V. corneae and E. bieneusi via intraperitoneal, oral or rectal route. 49.1.3.2  Molecular Techniques In contrast to conventional diagnostic procedures that use the phenotypic characteristics of microsporidia, which can be influenced by changing external conditions, molecular techniques have several distinct advantages. Firstly, as ­molecular techniques detect differences at the nucleotide level, they are much precise and not affected by variations resulting from

Molecular Detection of Foodborne Pathogens

altered temperature, osmolarity, pH, and nutrient availability, etc. Secondly, many current molecular detection formats such as PCR and its derivatives involve nucleic acid amplification, and they provide an unsurpassed sensitivity. Not surprisingly, molecular techniques have become the methods of choice for microsporidial species identification and strain/genotype determination.18 Although many genetic components in microsporidia can be targeted for diagnostic purposes, small (16S) and large (23S) subunit rRNA and α- and β-tubulin are favored due to the relative sequence conservation among these genes and the ready accessibility of their nucleotide sequences from various microsporidial species. Species-specific detection. Ribosomal RNA molecules (rRNAs) are key components of the ribosome that is responsible for protein synthesis in all living organisms. Consisting of two complex folded structures of differing sizes, rRNAs provide a mechanism for decoding messenger RNA (mRNA) into amino acids (at center of small ribosomal subunit) and to interact with the transfer RNA (tRNA) during translation by providing petidyltransferase activity (large subunit). Because of its critical role in cellular function and maintenance, rRNAs are not only the most conserved, but also the most abundant in all cells. Although being considered as eukaryotes, microsporidia harbor small (16S) and large (23S) subunit rRNA genes that resemble those found in prokaryotes, and they also lack a separate 5.8S rRNA that is characteristic of eukaryotes. The small (16S) and large (23S) subunit rRNA genes and their ITS regions has been successfully utilized for the species-specific detection and differentiation of microsporidia infecting humans and animals (e.g., E. bieneusi, E. cuniculi, E. hellem, E. intestinalis, T. hominis, V. corneae, V. necatrix).18,19–25 The tubulin gene family in eukaryotic organisms ­comprises three distinct, and yet conserved subfamilies (i.e., α-, β-, and γ-tubulin) that encode proteins essential for the formation of microtubules, which result from the polymerization of tubulin consisting of a dimer of α- and β-subunits, each about 440 amino acids long. Microtubules constitute the major components of the eukaryotic cytoskeleton and mitotic spindle. The α- and β-tubulins are abundant in eukaryotic cells, and their underlying genes are remarkably stable with their nucleotide deletions or insertions being rarely observed. These features make the α- and β-tubulin gene sequences valuable targets for identification and phylogenetic analysis of mitochondrial microsporidia and protozoa. Indeed, a number of reports have described the use of α-tubulin gene for molecular determination of Nosema locustae and Spraguea lophii.26 and β-tubulin gene for identification of E. cuniculi, E. hellem, and E. intestinalis.14,27 In addition, other genes have been also employed for species-specific detection and identification of microsporidia. These include genes encoding isoleucyl-tRNA synthase, glutamyl-tRNA synthetase, U2, and U6 spliceosomal RNA, reverse transcriptase, chitin synthase, and DNA-directed RNA polymerase II, translation elongation factors EF-1α and EF-2, a chaperone protein Hsp70, actin (ACT1), a polar tube protein, a ribosomal protein, dihydrofolate reductase, serine hydroxymethyltransferase, aminopeptidase, and thymidylate synthase.18

695

Encephalitozoon and Enterocytozoon

Genotyping. Of the two genera in the family Enterocytozoonidae, the genus Nucleospora contains two characterized species: (i) N. salmonis, an intranuclear microsporidia of salmonid fish, and (ii) N. secunda, a parasite of a warm-water African fish; while the genus Enterocytozoon includes E. bieneusi only, which infects the cytoplasm of enterocytes and other epithelial cells in humans and mammals and is the most common microsporidia species known to cause human disease. Although only one genus has been recognized in the family Encephalitozoonidae, recent studies indicate that E. hellem, E. cuniculi, and E. intestinalis may form three distinct genera as they show notable morphological and biological differences. For example, in contrast to E. cuniculi, some E. hellem parasites are located within a parasitophorous vacuole, while others are present in the host cell cytoplasm.28 Based on the β-tubulin sequences, it appears that E. intestinalis is possibly more closely related to E. cuniculi than to E. hellem.29 Susceptibility testing. Mutations in microsporidial β-tubulin gene have been shown to be linked to susceptibility or resistance to a number of antiparasitic drugs. Being effectors of microtubule polymerization, benzimidazoles (e.g., albendazole) disrupt microsporidial microtubules through β-tubulin binding, and some microsporidia induce mutations in His-6, Phe-167, Glu-198, Phe-200, and Arg-241 to enhance resistance to the derivate benomyl, and in Ala-165 to confer resistance to thiabendazole resistance.29 On the other hand, E. hellem and E. cuniculi β-tubulins with nonaltered His-6, Phe-167, Glu-198, Phe-200, and Arg-241 are generally susceptible to benzimidazoles.30



49.2  Methods











49.2.1 Sample Preparation Sample collection. Fresh specimens (e.g., stool, sputum, bronchoalveolar lavage fluid, nasal secretion, or cerebrospinal fluid, conjunctival smears, corneal scrapings, or tissue and body fluids) are preferred for molecular studies). If formalin-fixed fecal samples are used the formalin has to be removed by washing/centrifuging steps with saline direct or after formalin-ether concentration. However, sensitivity of amplification is known to decrease with time of fixation. DNA isolation. Both in-house and commercial reagents have been used for preparation of DNA from microsporidia. We present two methods below that are often used in our laboratory. (i) DNA isolation from feces using QIAamp Spin Columns*31: (1) Suspend 100 mg of feces in 200 µl of PBS with 2% PVPP (polyvinylpolypyrolidone, Sigma P6755).† This adaptation of the QIAamp DNA minikit cat. No. 51306 gives better results than QIAamp stool kit. † Or suspend 0.5 g of feces in 2 ml of PBS with 2% PVPP and seave (filter). For isolation of the DNA aliquot a separate amount of 200 µl in an Eppendorf tube, and store the rest at -70°C. PVPP does not dissolve; it stays like a suspension so you have to stir regularly while using it. *



‡

(2) Heat feces-suspension for 10 min in heatblock at 100°C. (3) Add an equal volume of tissue lysis buffer (ATL) containing 20 µl proteinase K (so that is 180 µl of ATL and 20 µl of prot K from the kit) solution to the sample (vortex). Incubate 2 h or overnight at 55°C in heatblock (after 1 h vortex briefly). (4) Add 400 µl AL buffer (with spike) to 400 µl sample, mix thoroughly by vortexing, and incubate at 70°C for 10 min. (5) Centrifuge 30 sec full speed and transfer supernatant to an Eppendorf containing 400 µl of ethanol (96–100%), and mix thoroughly by vortexing (short spin). (6) Place a numbered spin column in a 2 ml collection tube. Carefully apply 600 µl of the mixture from step 5 to the spin column without moistening the rim, close the cap and centrifuge at 10000 rpm for 1 min. (7) Place spin column in a clean 2 ml collection tube and repeat with the rest of the mixture from step 5. (8) Place the spin column in a clean 2 ml collection tube. (9) Carefully open the spin column and add 500 µl of AW1 buffer. Centrifuge at 10000 rpm for 1 min. (10) Empty the collection tube and add 500 µl of AW2 buffer. Centrifuge 3 min, 1 min at 10000 rpm and 2 min at full speed. (11) Place the spin column in a numbered, clean 1.5 ml microfuge tube (with lid). (12) Carefully open the spin column, elute the DNA with 200 µl AE buffer, leave the buffer on the filter for 1 min and centrifuge at 8000 rpm for 1 min. (ii) DNA isolation from feces using Magnapure: (1) Suspend 100 mg of feces in 200 µl of PBS with 2% PVPP.‡ (2) Heat feces-suspension for 10 min in heatblock at 100°C. (3) Add an equal volume of tissue lysis buffer (ATL) (Qiagen) containing 20 µl proteinase K (so that is 180 µl of ATL and 20 µl of prot K from the kit) solution to the sample (vortex). Incubate 2 h or overnight at 55°C in heatblock (after 1 h vortex briefly). (4) Centrifuge 30 sec full speed and transfer supernatant to the Magnapure sample-cartridge. (5) Perform isolation using the Magnapure bacterial III kit. (6) Add PhHV (to serve as an internal control) to the Magnapure Lysis buffer. (7) Elute DNA in 100 µl. Or suspend 0.5 g of feces in 2 ml of PBS with 2% pvpp (polyvinylpolypyrolidone) and filter. For isolation of the DNA, aliquot a separate amount of 200 µl in an Eppendorf tube and store the rest at –70°C.

696

Molecular Detection of Foodborne Pathogens

BSA

Table 49.1 Primers for Multiplex Nested PCR Oligo Name

Sequence

Primer Msp-1

5′-TGAATGKGTCCCTGT-3′

Primer Msp-2A

5′-TCACTCGCCGCTACT-3′

Primer Msp-2B

5′-GTTCATTCGCACTACT-3′

Primer Msp-3

5′-GGAATTCACACCGCCCGTCRYTAT-3′

Primer Msp-4A

5′-CCAAGCTTATGCTTAAGTCCAGGGAG-3′

Primer Msp-4B

5′-CCAAGCTTATGCTTAAGTYMAARGGGT-3′

49.2.2  Detection Procedures 49.2.2.1  Mutiplex Nested PCR20 Principle: A multiplex nested PCR protocol has been reported for differentiation of E. bieneusi from other microsporidia commonly found in humans.20 This PCR generates a 508-bp fragment of the SSU rRNA gene in the first round amplification with primers MSP-1 and MSP-2B, which consists of 122 bp of the 3′-end of the SSU rRNA gene, 243 bp of the ITS and 143 bp of the 5′-region of the LSU rRNA gene.20 The second round amplification is performed with primers MSP-3 and MSP-4B (Table 49.1). Procedure: (1) Prepare the first round PCR mixture (20 µl) as ­follows, and add 5 µl DNA in the test tube (or ­distilled water in the negative control tube): First Round PCR

1 × mix(µl)

20 × mix(µl)

Hotstart mastermix Primer Msp-1 Primer Msp-2A Primer Msp-2B BSA H2O Total

12.5 0.5 0.5 0.5 0.5 6.5 20.00

250 10 10 10 10 130 400

25 uM 25 uM 25 uM 5 mg/ml

(2) Run PCR amplification in a thermocycler for one cycle of 95ºC for 15 min; 25 cycles of 95ºC for 15 sec, 55ºC for 30 sec, and 72ºC for 30 sec; one cycle of 72ºC for 7 min. (3) Separate PCR products (10 µl) on 2% agarose gel containing ethidium bromide and visualize with UV transilluminator. (4) Prepare the second round PCR mixture (20 µl) as follows, and add 5 µl of PCR product from the first round PCR in each tube (or distilled water in the negative control tube). 1 × mix(µl)

Hotstart mastermix

0.5

10

6.5 20.00

130 400

(5) Perform PCR amplification and detect as above (steps 4 and 5). Note: This nested PCR has a detection limit of three to 100 spores in a 0.1-g stool sample, and allows discrimination among E. bieneusi, E. cuniculi and E. intestinalis upon sequencing analysis. Indeed, along with the primer pair EBIEF1-EBIER1 (i.e., 5′-GAAACTTGTCCACTCCTTACG-3′ and 5′-CCATGCACCACTCCTGCCATT-3′, permitting amplification of a 607-bp fragment from SSU rRNA gene of E. bieneusi),32 the primer pair MSP3-MSP4B has been shown to offer one of the most sensitive methods for detection of E. bieneusi from stools.33 49.2.2.2 Real-time PCR25 Principle: A multiplex real-time PCR method was developed for the simultaneous detection of Enterocytozoon ­bieneusi and Encephalitozoon spp. in stool samples.25 The E. bieneusi-specific real-time PCR amplifies and detects a 103-bp fragment within the ITS sequence of E. bieneusi. The Encephalitozoon spp. PCR primers and detection probe were designed on the SSU rRNA gene sequence for E. intestinalis, amplifying, and detecting a 214-bp fragment inside the SSU rRNA gene. This multiplex PCR included an internal control to detect inhibition of the amplification by fecal constituents in the sample (Table 49.2). Procedure: (1) Prepare PCR mixture (25 μl) containing PCR buffer (Hotstart Taq master mix, QIAgen), 5 mmol/l MgCl2, 2 pmol of each E. bieneusi-specific primer, 6.25 pmol of each E. intestinalis-specific primer, 3.75 pmol of each PhHV-1-specific primer, 2.5 pmol of E. bieneusi-specific double-labeled probe, 2.5 pmol of Encephalitozoon spp.-specific doublelabeled probe, 2.5 pmol of PhHV-1–specific double-labeled probe, and 5 μl of the DNA sample as follows: 1 × mix (µl) Hotstart mastermix

12.50

100 × mix (µl) 1250

primer Eb F–EbITS-89F

10 uM

0.20

20.00

primer Eb R–EbITS-191R

10 uM

0.20

20.00

10 uM

0.25

25

probe Eb–EbITS-114revT Second Round PCR

5 mg/ml

H2O Total

FAM

20 × mix(µl)

primer MSP–MSP1F

25 uM

0.25

25.00

12.5

250

primer R Ei–Ein227R

25 uM

0.25

25.00

VIC

Primer Msp-3

25 uM

0.5

10

probe Ei–Enc82trev

10 uM

0.25

25

Primer Msp-4A

25 uM

0.5

10

primer PhHV F–267S

25 uM

0.15

15.00

Primer Msp-4B

25 uM

0.5

10

primer PhHV R–337 AS

25 uM

0.15

15.00

697

Encephalitozoon and Enterocytozoon

probe PhHV–305Tq MgCl2

CY5

10 uM

0.25

25 mM

3.50

350.00

0.80

80.00

0.50

50

0.75

75.00

Rox 1:20* BSA H2O

5 mg/ml

25

Total 20.00 2000.00 *Only needed for ABI 7500 can be left out for BioRad Icycler.

(2) Perform PCR in I-cycler real-time detection system (BioRad, Hercules, CA) for one cycle of 95°C for 15 min; 50 cycles of 95°C for 15 sec, 60°C for 30 sec, and 72°C for 30 sec. (For amplification in ABI 7500, the cycling program consists of one cycle of 95ºC for 15 min; 50 cycles of 95ºC for 15 sec, and 60ºC for 60 sec). (3) Detect fluorescence of FAM, Yakima Yellow and CY5 at their respective wavelengths during the annealing step of each cycle. Note: The Encephalitozoon spp. primers and probe were designed on the known sequence of E. intestinalis. According to NCBI BLAST, however, E. hellem and E. cuniculi will be amplified and detected also with this primer/probe combination. Although determination to the genus level will be sufficient in clinical practice, species-specific differentiation can be achieved by sequencing of the PCR product. This PCR was evaluated with species-specific DNA controls (n = 22) and a range of well-defined stool samples (n = 140), and it demonstrated 100% specificity and sensitivity.25 Thus, this assay offers the possibility of introducing

DNA detection as a feasible technique in the routine diagnosis of intestinal microsporidia infections. 49.2.2.3 Genotyping of Enterocytozoon bieneusi on the ITS Region Principle: This PCR amplifies a 296-bp fragment of the ITS-1 region. After purification of the PCR product, both the forward and reverse strands are sequenced with the same primers used for initial PCR.44 Oligo Name

Sequence

Eb-80F

5′-GTTGGAGAACCAGCTGAAGGAT-3′

Eb-375R

5′-ATACACCTCTTGATGGCACCCT-3′

Procedure: (1) Prepare PCR mixture (20 μl) as follows, and add 5 µl DNA in the test tube (or distilled water in the negative control tube):

Hot start master mix Eb-80F Eb-375R BSA H2O Total

25 µM 25 µM 5 mg/ml

1×

20

12.5 0.5 0.5 0.5 6 20

250 10 10 10 120 400

(2) Perform PCR amplification for 1 cycle of 95ºC for 15 min; 25 cycles of 95ºC for 15 sec, 55ºC for 30 sec, and 72ºC for 30 sec; 1 cycle of 72ºC for 7 min.

Table 49.2 Oligonucleotide Primers and Detection Probes for Real-Time PCR for the Simultaneous Detection of Enterocytozoon bieneusi, Encephalitozoon spec, and Phocin Herpes Virus 1 as an Internal Control Target Organism Oligonucleotide Name

Oligonucleotide Sequence

Genbank Accession no. or Literature Reference of Target Sequence

Enterocytozoon bieneusi EbITS-89F

5′-TGTGTAGGCGTGAGAGTGTATCTG-3′

AF101198

EbITS-191R

5′-CATCCAACCATCACGTACCAATC-3′

AF101198

EbITS-114revT

FAM-5′-CACTGCACCCACATCCCTCACCCTT-3′-Eclipse

AF101198

MSP1F

5′-CACCAGGTTGATTCTGCCTGAC-3′

U09929

Eint227R

5′-CTAGTTAGGCCATTACCCTAACTACCA-3′

U09929

Eint82Trev

Yakima Yellow-5′-CTATCACTGAGCCGTCC-3′-Eclipse

U09929

PhHV-267s

5′-GGGCGAATCACAGATTGAATC-3′

Niesters, 200234

PhHV-337as

5′-GCGGTTCCAAACGTACCAA-3′

Niesters, 200234

PhHV-305tq

Cy5 5′-TTTTTATGTGTCCGCCACCATCTGGATC-3′-BHQ2

Niesters, 200234

Encephalitozoon spec.

Phocin Herpes Virus 1

698

Molecular Detection of Foodborne Pathogens

(3) Separate PCR products (10 µl) on 2% agarose gel containing ethidium bromide and visualize with UV transilluminator. (4) Purify PCR products using for example the High Pure PCR Product Purification Kit (Roche Diagnostics, Mannheim, Germany) according to the manufacturer’s protocol. (5) Measure DNA concentration with for example Nanodrop ND-1000 UV-Vis Spectrophotometer (Nanodrop Technologies, DE). (6) Perform sequcing reaction for both the forward and reverse strands with the same primers used for PCR.



Note: A large number of primers have been designed from E. bieneusi ITS sequence and utilized for identification and genotyping purposes. Some of these primers may assume different names in the literature, but possess identical nucleotide sequences.







49.3 Conclusion and Future Perspectives As microsporidial genera and species demonstrate biological and epidemiological variations which often demand different approaches to prevention and treatment, it is necessary to accurately determine their taxonomic status. The microscopic diagnosis of microsporidia infections using additional staining techniques is labor-intensive, therefore this will often be limited to samples of severe immune-suppressed patients. Many laboratories will not see as many of these patients due to the success of HAART therapy in HIV infected individuals. As a consequence gaining expertise is difficult as positive findings will be scarce. Moreover, in recent years intestinal microsporidiosis is increasingly diagnosed in other immunocompromised individuals such as organ transplant recipients, as well as in children, travelers, and elderly.13,35–43 Nucleic acid-based amplification methods for the detection of microsporidia give useful alternatives for the labor intensive and expertise depended microscopy. Especially realtime PCR, with its reduced risk for contamination and the possibility of multiplex detection of different targets in one tube, offers the possibility of introducing DNA detection as a feasible technique in the routine diagnosis of intestinal microsporidial infections and can also be used as a part of a series of other multiplex assays in the differential diagnosis of diarrhea causing pathogens.

References

1. Vossbrinck, C.R. et al. Ribosomal RNA sequence suggests microsporidia are extremely ancient eukaryotes. Nature, 326, 411, 1987. 2. Vivares, C.P. et al. Functional and evolutionary analysis of a eukaryotic parasitic genome. Curr. Opin. Microbiol., 5, 499, 2002.









3. Keeling, P.J. and Fast, N.M. Microsporidia: Biology and evolution of highly reduced intracellular parasites. Annu. Rev. Microbiol., 56, 93, 2002. 4. Gill, E.E. and Fast, N.M. Assessing the microsporidia-fungi relationship: Combined phylogenetic analysis of eight genes. Gene, 375, 103, 2006. 5. Sprague, V., Becnel, J.J. and Hazard, E.I. Taxonomy of phylum microspora. Crit. Rev. Microbiol., 18, 285, 1992. 6. Didier, E.S. Microsporidiosis: an emerging and opportunistic infection in humans and animals. Acta Trop., 94, 61, 2005. 7. Didier, E.S. and Weiss, L.M. Microsporidiosis: current status. Curr. Opin. Infect. Dis.,19, 485, 2006. 8. Didier, E.S. et al. Epidemiology of microsporidiosis: sources and modes of transmission. Vet. Parasitol., 126, 145, 2004. 9. Franzen, C. How do microsporidia invade cells? Folia Parasitol. (Praha), 52, 36, 2005. 10. Franzen, C. and Salzberger, B. Analysis of the beta-tubulin gene from Vittaforma corneae suggests benzimidazole resistance. Antimicrob. Agents Chemother., 52, 790, 2008. 11. Sulaiman, I.M. et al. A molecular biologic study of Enterocytozoon bieneusi in HIV-infected patients in Lima, Peru. J. Eukaryot. Microbiol., 50, 591, 2003. 12. Didier, E.S. et al. Identification and characterization of three Encephalitozoon cuniculi strains. Parasitology, 111, 411, 1995. 13. Gomez Morales, M.A. et al. Opportunistic and non-­opportunistic parasites in HIV-positive and negative patients with diarrhoea in Tanzania. Trop. Med. Parasitol., 46, 109, 1995. 14. Mathis, A., Weber, R. and Deplazes, P. Zoonotic potential of the microsporidia. Clin. Microbiol. Rev., 18, 423, 2005. 15. Sulaiman, I.M. et al. Molecular characterization of Enterocytozoon bieneusi in cattle indicates that only some isolates have zoonotic potential, Parasitol. Res., 92, 328, 2004. 16. van Gool,T., Canning, E.U. and Dankert, J. An improved practical and sensitive technique for the detection of microsporidian spores in stool samples. Trans. R. Soc. Trop. Med. Hyg., 88, 189, 1994. 17. Garcia, L.S. Laboratory identification of the microsporidia. J. Clin. Microbiol., 40, 1892, 2002. 18. Franzen, C. Microsporidia: how can they invade other cells? Trends Parasitol., 20, 275, 2004. 19. Zhu, X. et al. Small subunit rRNA sequence of Enterocytozoon bieneusi and its potential diagnostic role with use of the polymerase chain reaction. J. Infect. Dis., 168, 1570, 1993. 20. Katzwinkel-Wladarsch, S. et al. Direct amplification and species determination of microsporidian DNA from stool specimens. Trop. Med. Int. Health, 1, 373, 1996. 21. Wolk, D.M. et al. Real-time PCR method for detection of Encephalitozoon intestinalis from stool specimens. J. Clin. Microbiol., 40, 3922, 2002. 22. Menotti, J. et al. Development of a real-time polymerase-­ chain-reaction assay for quantitative detection of Enterocytozoon bieneusi DNA in stool specimens from immunocompromised patients with intestinal microsporidiosis. J. Infect. Dis., 187, 1469, 2003. 23. Menotti, J. et al. Development of a real-time PCR assay for quantitative detection of Encephalitozoon intestinalis DNA. J. Clin. Microbiol., 41, 1410, 2003. 24. Notermans, D.W. et al. Detection and identification of Enterocytozoon bieneusi and Encephalitozoon species in stool and urine specimens by PCR and differential hybridization. J. Clin. Microbiol., 43, 610, 2005.

Encephalitozoon and Enterocytozoon 25. Verweij, J.J. et al. Multiplex detection of Enterocytozoon ­bieneusi and Encephalitozoon spp. in fecal samples using real-time PCR. Diagn. Microbiol. Infect. Dis., 57, 163, 2007. 26. Mathis, A. et al. Two Encephalitozoon cuniculi strains of human origin are infectious to rabbits. Parasitology, 114, 29, 1997. 27. Mathis, A. et al. Genetic and phenotypic intraspecific variation in the microsporidian Encephalitozoon hellem. Int. J. Parasitol., 29, 767, 1999. 28. Cali, A., Weiss, L.M., and Takvorian, P.M. Microsporidian taxonomy and the status of Septata intestinalis. J. Eukaryot. Microbiol., 43, 106S, 1996. 29. Li, J. et al. Tubulin genes from AIDS-associated microsporidia and implications for phylogeny and benzimidazole sensitivity. Mol. Biochem. Parasitol., 78, 289, 1998. 30. Katiyar, S.K. et al. Antiprotozoal activities of benzimidazoles and correlations with beta-tubulin sequence. Antimicrob. Agents Chemother., 38, 2086, 1994. 31. Verweij, J.J. et al. Determining the prevalence of Oesophagostomum bifurcum and Necator americanus infections using specific PCR amplification of DNA from faecal samples. Trop. Med. Int. Health, 6, 726, 2001. 32. da Silva, A.J. et al. Sensitive PCR diagnosis of infections by Enterocytozoon bieneusi (microsporidia) using primers based on the region coding for small-subunit rRNA. J. Clin. Microbiol., 34, 986, 1996. 33. Subrungruang, I. et al. Evaluation of DNA extraction and PCR methods for detection of Enterocytozoon bienuesi in stool specimens. J. Clin. Microbiol., 42, 3490, 2004. 34. Niesters, H.G. Clinical virology in real time. J. Clin. Virol., 25 (suppl), S3, 2002.

699 35. Wanke, C.A., DeGirolami, P. and Federman, M. Enterocytozoon bieneusi infection and diarrheal disease in patients who were not infected with human immunodeficiency virus: case report and review. Clin. Infect. Dis., 23, 816, 1996. 36. Gainzarain, J.C. et al. Detection of Enterocytozoon bieneusi in two human immunodeficiency virus-negative patients with chronic diarrhea by polymerase chain reaction in duodenal biopsy specimens and review. Clin. Infect. Dis., 27, 394, 1998. 37. Raynaud, L. et al. Identification of Encephalitozoon ­intestinalis in travelers with chronic diarrhea by specific PCR amplification. J. Clin. Microbiol., 36, 37, 1998. 38. Lopez-Velez, R. et al. Microsporidiosis in travelers with diarrhea from the tropics. J. Travel Med., 6, 223, 1999. 39. Muller, A. et al. Detection of microsporidia in travelers with diarrhea. J. Clin. Microbiol., 39, 1630, 2001. 40. Lores, B. et al. Intestinal microsporidiosis due to Enterocytozoon bieneusi in elderly human immunodeficiency virus--negative patients from Vigo, Spain. Clin. Infect. Dis., 34, 918, 2002. 41. Leelayoova, S. et al. Transmission of Enterocytozoon bieneusi genotype a in a Thai orphanage. Am. J.Trop. Med. Hyg., 73, 104, 2005. 42. Mungthin, M. et al. Spore shedding pattern of Enterocytozoon bieneusi in asymptomatic children. J. Med. Microbiol., 54, 473, 2005. 43. Wichro, E. et al. Microsporidiosis in travel-associated ­chronic diarrhea in immune-competent patients. Am. J. Trop. Med. Hyg., 73, 285, 2005. 44. ten Hove, R.-J. et al. Characterization of Enterocytozoon ­bieneusi genotypes in immunosuppressed and immunocompetent individuals, (manuscript submitted).

50 Giardia Yaoyu Feng

East China University of Science and Technology

Lihua Xiao

Centers for Disease Control and Prevention

Contents 50.1 Introduction.................................................................................................................................................................... 701 50.1.1 Classification of Giardia................................................................................................................................... 702 50.1.2 Biology, Pathogenesis, and Medical Importance . ........................................................................................... 702 50.1.2.1 Life Cycle......................................................................................................................................... 702 50.1.2.2 Transmission.................................................................................................................................... 702 50.1.2.3 Epidemiology................................................................................................................................... 702 50.1.2.4 Pathogenesis..................................................................................................................................... 703 50.1.3 A Review of Methods for Diagnosis of G. duodenalis..................................................................................... 703 50.1.3.1 Conventional Diagnosis .................................................................................................................. 703 50.1.3.2 Molecular Methods Available and their Characteristics/Performance............................................ 704 50.2 Methods...........................................................................................................................................................................710 50.2.1 Reagents and Equipment....................................................................................................................................710 50.2.1.1 Supplies for Sample Collection and Preservation.............................................................................710 50.2.1.2 Supplies for Filtration and Centrifugation........................................................................................710 50.2.1.3 Supplies for IMS...............................................................................................................................710 50.2.1.4 Supplies for DNA Extraction............................................................................................................710 50.2.1.5 Supplies for PCR...............................................................................................................................710 50.2.1.6 Supplies for DNA Sequencing..........................................................................................................710 50.2.2 Sample Preparation............................................................................................................................................710 50.2.2.1 Fecal Sample Collection and Preservation.......................................................................................710 50.2.2.2 Extraction of Giardia Cysts from Food and Water Samples............................................................710 50.2.2.3 Centrifugation of Giardia Cysts from Food and Water Samples.....................................................711 50.2.2.4 Purification of Giardia Cysts from Food and Water Samples..........................................................711 50.2.2.5 DNA Extraction from Fecal Specimens, Food, and Water Samples................................................711 50.2.3 Detection Procedures........................................................................................................................................ 712 50.2.3.1 Primary PCR................................................................................................................................... 712 50.2.3.2 Secondary PCR................................................................................................................................ 712 50.2.3.3 Detection of Secondary PCR Products . ......................................................................................... 712 50.2.3.4 DNA Sequencing............................................................................................................................. 712 50.2.3.5 Sequence Analysis........................................................................................................................... 712 50.3 Conclusions and Further Perspective ............................................................................................................................ 713 Acknowledgments .................................................................................................................................................................... 713 References.................................................................................................................................................................................. 713

50.1  Introduction Giardia is a genus of protozoa that infect a wide range of vertebrates. One species within this genus, Giardia duodenalis (syn. G. lamblia and G. intestinalis), causes human giardiasis, which probably constitutes the most common cause of protozoan diarrhea worldwide, leading to significant morbidity and mortality in both the developing and developed world.

Giardiasis is one of the major causes of waterborne and foodborne outbreaks of diarrhea in industrialized nations.1 Waterborne outbreaks of giardiasis were first documented in Aspen, CO, in 1965/1966, and the Centers for Disease Control and Prevention (CDC) began waterborne disease surveillance in 1971. Between 1979 and 1988, Giardia was the most frequently implicated organism in waterborne disease 701

702

outbreaks. Between 1965 and 1984, some 90 outbreaks with a total of 23,776 cases were reported in the United States.2 Between 1992 and 1997, national surveillance carried out by 43 states in the United States indicated that as many as 2.5 million cases of giardiasis occur annually in the country.3 Waterborne and foodborne giardiasis is well known among travelers to countries in Eastern Europe and the former Soviet Union.4 A number of outbreaks of foodborne giardiasis related to food preparation have been documented, probably caused by infected food handlers or contact by food handlers with infected people, particularly children.4 For example, an outbreak of giardiasis occurred after a family party for 25 people. Nine people who ate fruit salad at the party became ill. The person who prepared the fruit salad had a diapered child and a pet rabbit at home who were both positive for G. duodenalis.5 In 1990, an outbreak among insurance company employees resulted in 18 laboratory-confirmed and nine suspected cases of giardiasis. Raw sliced vegetables served in the employee cafeteria and prepared by a food handler infected with G. duodenalis were the probable cause of the outbreak.6 No outbreaks of foodborne giardiasis related to industrially manufactured foods have been reported.

50.1.1 Classification of Giardia The nomenclature for Giardia species is confusing. Giardia species are established based on morphology and host specificity. Six species are generally recognized, including G. agilis in amphibians, G. ardeae and G. psittaci in birds, G. muris in rodents, G. microti in muskrats and voles, and G. duodenalis in mammals. Humans are believed to be infected by a single species, variably termed G. lamblia, G. intestinalis, or G. duodenalis. For consistency, G. duodenalis is used in this chapter. Within this species the current trend has been to identify a complex of assemblages based on genetic differences and host specificity. These assemblages are identified based on sequence analysis of conserved genetic loci.7 Currently, there are seven well defined assemblages of G. duodenalis, designated A through G. Assemblages A and B have the broadest host specificity, having been found to infect humans and other mammals, including dogs, cats, livestock, and wildlife.7 Assemblage A consists of mostly two subgroups, AI, and AII, but there are no clear subgroups in assemblage B.8–13 Assemblages C and D have been found only in dogs.14–16 Assemblage E has been found only in clovenhoofed mammals.17 Assemblages F and G have been found in cats and rodents, respectively.8

50.1.2 Biology, Pathogenesis, and Medical Importance 50.1.2.1 Life Cycle Giardia has a simple and direct life cycle. Cysts excreted in feces contain a mitotically arrested trophozoite that can remain infectious for months in a wet, cool environment.

Molecular Detection of Foodborne Pathogens

After cysts are ingested, trophozoites emerge in the duodenum and attach to the surface of the intestinal microvilli beneath the mucus layer via a ventral adhesive disc. Trophozoites undergo repeated mitotic division. Environmentally resistant cysts form from trophozoites in response to stimuli of intestinal conditions such as the presence of major bile salts. They pass through the intestine in the feces and are spread by contaminated water, food, and fomites, and by physical contact. Infection of new hosts is established by ingestion of cysts. 50.1.2.2 Transmission Giardiasis is transmitted via the fecal–oral route following direct or indirect contact with Giardia cysts, including person to person, zoonotic, waterborne, and foodborne transmission.18 In industrialized nations, Giardia cysts are frequently transmitted through drinking and recreational water. The contributors to environmental contamination include infected human, livestock, and feral hosts as well as transport hosts. Cysts in feces can contaminate water directly or indirectly, and the disposal of human and animal wastes remains a significant public health issue in many parts of the world. Transmission of Giardia through drinking and recreational water is well documented, as there have been numerous outbreaks of giardiasis following the ingestion of contaminated water.19,20 Foods that have been associated with cyst contamination from human or animal sources include raw vegetables and fruit salad. Fresh produce consumed with minimal preparation, in particular, is a potential vehicle of transmission, and G. duodenalis cysts have been detected on produce in several countries.21–24 Contaminated irrigation water appears be a major route of contamination of fresh produce.25,26 There are also reports of cyst contamination of herbs,21,27,28 surface waters used for salad crop irrigation, and wash water at packing houses.25 Although most documented foodborne outbreaks of giardiasis were attributed to direct contamination by food handlers,5,6,29–31 reports of two outbreaks implicated zoonotic transmission as the causes; one was the consumption of a Christmas pudding contaminated with rodent feces32 whereas the other was the consumption of a tripe soup made from the offal of an infected sheep.33 50.1.2.3 Epidemiology Estimates of prevalence for giardiasis vary greatly because reporting is not universally required, diagnostic methods vary greatly, and many infected persons in endemic areas are not symptomatic, have no access to medical care, or do not seek medical treatment. In the United States, G. ­duodenalis is among the most commonly diagnosed parasitic enteric pathogens. Giardia duodenalis has been identified in the United States in 4–7% of stool specimens from patients who have diarrheal illness. Voluntary reporting to the CDC’s Nationally Notifiable Disease Surveillance System from 1998 to 2005 indicated that the total number of cases varied between 19,708 and 24,226 annually.34,35 However, because of underreporting, estimates of annual giardiasis cases are likely to be four to 100 times that of reported cases.36

703

Giardia

Giardiasis affects persons of all ages and peaks during the late summer and fall months. In the United States, the greatest number of cases of giardiasis was reported in children of 1–9 years of age and adults of 30–39 years of age.35 A multicenter study of outpatient pediatric nondysenteric diarrhea in the United States found that Giardia was the cause of diarrhea in 15% of the children.37 In developing countries, Giardia is one of the first enteric pathogens to infect infants, with a prevalence rate up to 30% in children who are younger than 10 years of age.38–42 Giardia has been one of the most common pathogens in outbreaks of diarrheal illness associated with drinking and recreational water.43 Recreational, surface, and well water can contain Giardia cysts. A case control study in southwestern England showed that swallowing water while swimming, contact with recreational fresh water, drinking tap water, and eating lettuce were risk factors for developing giardiasis.44 The association with eating raw lettuce (and other fruit and vegetables which receive minimal heat treatment before consumption) highlights the role of contaminated wastewater, uncomposted sewage sludge or manure used as fertilizer, in addition to direct fecal contamination of produce by farm animals and wildlife, or transmission vectors such as refuse-feeding birds and non-biting filth flies. In the United States, a seasonal peak in age-related giardiasis cases occurs in the summer recreational water season, possibly reflecting increased use of rivers, lakes, swimming pools, and water parks.35 Person-to-person transmission occurs in groups who have poor fecal-oral hygiene. Among children in day care centers, Giardia cyst passage was documented to be as high as 50%.45 Many of these infected children are asymptomatic and can spread the cysts among household members. The Giardia cyst passage rate is as high as 20% among sexually active gay men.46 Giardia cysts were reported to be transmitted in commercial food establishments, corporate office settings, small gatherings, and travelers to developing countries.6,47,48 Molecular tools have enabled not only the identification of species and genotypes in the feces of infected hosts but also the tracking of contamination sources in the environment, including water and fresh produce. However, less progress has been made in elucidating the epidemiology of giardiasis. Only G. duodenalis assemblages A and B have so far been associated with human infections. Because both assemblages A and B have also been found to infect animals, zoonotic transmission has long been suspected to play a role in epidemiology of human giardiasis. Although assemblage A has been considered the most important G. duodenalis genotype involved in zoonotic giardiasis, subtyping data do not support widespread findings of zoonotic transmission. The two most common subtypes of assemblage A, AI, and AII, differ significantly in host preference. Humans are mostly infected with AII, whereas animals are mostly infected with AI.49 50.1.2.4  Pathogenesis The pathogenesis of Giardia has been postulated to involve direct damage to the intestinal brush border and mucosa

by trophozoites, induction of a host immune response that results in the secretion of fluid and damage to the gut, alteration of bile content or duodenal flora, and apoptosis in the small intestinal epithelial cells.36 Disruption of the intestinal epithelial cell brush border may explain the lactose intolerance that commonly develops. In vitro studies showed that Giardia disrupts tight junctional zona occludens, increases permeability, and induces apoptosis in small intestinal epithelial cells.50 Increased permeability of the small intestinal epithelial cells occurs by way of a myosin light chain kinase-dependent phosphorylation of F-actin.51,52 However, mucosal invasion occurs rarely and no Giardia enterotoxin has been found. Giardia is capable of undergoing antigenic variations in its cysteine-rich surface proteins through a palmitoylation process during human infection and the encystation and excystation cycle.53,54 Antigenic variation protects the parasite from the activity of intestinal proteases, provides oxygen stability, and possibly allows adaptation to different hosts.36 The clinical significance of Giardia antigenic variation is not known.

50.1.3 A Review of Methods for Diagnosis of G. duodenalis 50.1.3.1  Conventional Diagnosis Traditionally, the diagnosis of Giardia infection has depended on microscopic examination of stool. This requires an experienced microscopist and a specialized laboratory. Stool samples should be examined fresh or placed immediately in a preservative. Motile trophozoites may be seen in a saline wet mount of fresh liquid stool from a patient who has acute diarrheal illness. Cysts can be identified after iodine staining or after preservation in 10% buffered formalin or polyvinyl alcohol and subsequent trichrome or iron hematoxylin staining. Repeated stool samples may be necessary to increase the sensitivity of diagnosis. The yield of cysts may be increased with formalin-ether or zinc sulfate flotation concentration techniques.55,56 Nevertheless, even under ideal conditions, organisms are identified in a single stool sample in only half of cases.36 Giardia stool antigen detection by enzymatic immunoassays (EIA) and immunochromatography (e.g., ImmunoCard STAT! Cryptosporidium/Giardia Rapid Assay, Meridian Bioscience, Inc.) is sometimes used in industrialized nations. However, direct fluorescence antibody (DFA) detection of cysts has become a common diagnostic practice for giardiasis in the United States. Commercial antigenic detection tests are similar in cost to microscopy with better reproducibility and more rapid return of results.57,58 Antigen tests have a sensitivity of 85–98% and a specificity of 90–100% compared with microscopy.57–61 A commonly used assay, ProSpecT (Giardia Microplate Assay, Remel, Lenexa, KS) detects cyst wall protein (CWP-1) by EIA, and Merifluor DFA (Meridian Bioscience, Inc.) uses a fluorescein-tagged monoclonal antibody against Giardia. The Merifluor DFA has a reported sensitivity and specificity of 96–100%,58,59 and is considered the gold standard by some diagnosticians.

704

Other tests that may be used for the diagnosis of Giardia include the string test or EnteroTest (HDC Corporation, Milpitas, CA), duodenal aspiration, duodenal biopsy, and serology. A string test that yields bile-stained mucus from the duodenum can be examined by wet mount for trophozoites. Although more invasive, duodenal aspiration and biopsy offer the advantage of examining for trophozoites and other enteric pathogens. For detection of Giardia in water, standardized methods, such as the U.S. Environmental Protection Agency Method 1623 and its equivalent in other countries,62–64 based on cyst entrapment by filtration, elution, and concentration of cysts by immunomagnetic separation (IMS), and identification by immunofluorescence and differential interference contrast microscopy using specified morphological, morphometric, and fluorescence criteria, can be used. These have provided valuable data on cyst occurrence. In contrast, there are no national or international guidelines for determining Giardia cyst contamination in or on foodstuffs and related studies are very limited. A method for detecting G. duodenalis cysts on lettuce can recover 46.0 ± 19.0% of G. duodenalis seeded (n = 30).65 This method was tested on a variety of commercially available natural foods and can recover 36.5 ± 14.3% of G. duodenalis seeded (n = 20). The basic steps for this method are similar to those for water samples except that an extraction procedure is used to replace the filtration and elution for water samples. However, the above methods are difficult or impossible to distinguish cysts of different Giardia species and the assemblages of G. duodenalis as cyst morphometry cannot be used to ascribe Giardia species or genotypes. Therefore, they should be used together with appropriate molecular typing tools to provide insight into the species and assemblages that occur in water and food samples. Molecular tools also have the advantage in their potential for the assessment of contamination source and the human-infective potential of Giardia cysts in samples. Molecular methods usually also have higher detection sensitivities than microscopy, which makes them applicable for detecting low numbers of Giardia cysts in water and food samples. 50.1.3.2 Molecular Methods Available and their Characteristics/Performance Molecular biology has provided powerful new tools for characterizing Giardia, and the analysis of previously unrecognized genetic differences within this genus has revolutionized our understanding of both the taxonomy and epidemiology of human giardiasis. Although some simple PCR assays have been used in the detection of Giardia in clinical and environmental samples, more recent molecular tools are developed to differentiate Giardia at the species/assemblage and subtype levels. These tools are widely used in the identification of G. duodenalis genotypes and subtypes in clinical specimens.8,14,66–71 Variations between the methods lie in the gene(s) utilized (such as the small subunit rRNA (SSU rRNA), glutamate dehydrogenase (GDH), triose phosphate

Molecular Detection of Foodborne Pathogens

isomerase (TPI), elongation factor 1 alpha (EF1a), and beta giardin (β-giardin), variant surface protein (VSP)), the specificity of the assay (Giardia specific, G. duodenalis specific, or assemblage specific), and downstream procedures (restriction fragment length polymorphisms or RFLP, and or DNA sequencing of PCR products, or hybridization). The usage of these loci in genotyping and subtyping of G. duodenalis and their sequence characteristics have been reviewed recently.13 Some published molecular targets for species, genotype, and subtype differentiation of Giardia in animal, human specimens, and water samples are listed in Table 50.1. Among all the genes studied, SSU rRNA, GDH, TPI and β-giardin are used more frequently than other loci. Since different Giardia loci differ in total, synonymous, and nonsynonymous substitution rates, their applications vary from species differentiation, genotyping (assemblagedifferentiation), to subtyping. For example, substitution rates for the partial SSU rRNA, GDH, and TPI genes were reported to be 0.01, 0.06 and 0.12 substitutions per nucleotide, respectively.13 Thus, the conserved SSU rRNA gene is traditionally used for species and assemblage differentiation (mostly genotyping),14,68,72–74 whereas the most variable locus, TPI, is frequently used for subtyping clinical samples75,76 and the GDH locus, with a substitution rate midway between them, has a broad application spectrum.8,70 Although β-giardin has a low substitution rate (0.03 substitutions per nucleotide), it has demonstrated prominent subgrouping within some common G. duodenalis assemblages, and therefore, has also been used for determining and differentiating the subtypes within assemblages A, B, and E.13 However, β-giardin has yet to demonstrate consistent subtyping with respect to the other loci. Genotyping tools. The genotyping primers for SSU rRNA gene locus of Giardia from Weiss et al.,66 van Keulen et al.77 or Hopkins et al.14 amplify only the first fifth or the last tenth of the gene (1400 bp in total), and the products produced by these primer sets are too small to genotype all of the current assemblages. Notably, assemblage F (of cats) is identical to assemblage A until base 499.13 This has led to some inconclusive genotyping and potentially inaccurate results being reported. In the study by Berrilli et al.78 the cat isolate was genotyped as assemblage A using the 292-bp product primers of Hopkins et al.14 and in a study by Fayer et al.79 the cat isolates were genotyped as assemblage F using the same primers. This discrepancy may be due to an unsubstantiated substitution at the 5′ end of the SSU rRNA gene and the limited sequences available in GenBank.13 In contrast, the other loci are comparatively useful for genotyping usage. The coverage of the genes also varies. The sequence of the GDH gene covers at least the first 40% to 60% of the gene, and the TPI and β-giardin sequences cover 60% of the length of the genes by most primers. Therefore, these lengths are sufficient for genotyping samples to the assemblage and possibly subassemblage level. Within a gene, regions of different substitution rates may also be targeted for different applications. For example, the

GDH

GDH

TPI

TPI

TPI

TPI

TPI

TPI

TPI

TPI

Gene

GDH-F2: GAGAGGATCCTTGARCCNGAGCGCG GDH-R2: ACCTTCTAGAANCCNGCDATGTTNGC-GCC

TPI-F1: CCAGAATGTGTACCTAGAGGGG TPI-R1: TACTCTCCGAGCTCCTTCTGC TPI-F2: ATGCCTGCTCGTCGCCCCTTC TPI-R2: CACTGGCCAAGCT′ICTCGCAG TPI-F3: GGAGACCGACGAGCAAAGC TPI-R3: CTTGCCAAGCGCCTCAA TPI-F4: AATAGCAGCACARAACGTGTATCTG TPI-R4: CCCATGTCCAGCAGCATCT TPI-F5: AAATIATGCCTGCTCGTCG TPI-R5: CAAACCTTITCCGCAAACC TPI-F6: CCCTTCATCGGIGGTAACTT TPI-R6: GTGGCCACCACICCCGTGCC TPI-F7: ATYAAGAGCCACGTRGCGKC TPI-R7: CCATGATTCTRCGYCTTTCAG TPI-F8: CGAGACAAGTGTTGAGATGC TPI-R8: GGTCAAGAGCTTACAACACG TPI-F9: CCAAGAAGGCTAAGCGTGC TPI-R9: GCCACATGCCTATGTACGGG TPI-F10: GTTGCTCCCTCCTTTGTGC TPI-R10: GGCCTTGCGTTCATCCAGG TPI-F11: GCACAGAACGTGTATCTGG TPI-R11: CTCTGCTCATTGGTCTCGC TPI-F12: ATCGGYGGTAAYTTYAARTG TPI-R12: CACTGGCCAAGYTTYTCRCA TPI-F13: CCCTTCATCGGYGGTAAC TPI-R13: CCCGTGCCRATRGACCACAC TPI-R14: ACRTGGACYTCCTCYGCYTGCTC GDH-F1: TCCACCCCTCTGTCAACCTTTC GDH-R1: AATGTCGCCAGCAGGAACG

Primer (5′–3′)

1170

220

520

141

210

452

576

253

532

605

81

148

683

812

Size (bp)

Could amplify most Giardia species and some bacteria

Genus-specific

Unknown

Assemblage B specific

Assemblage A specific

Unknown

Assemblage A specific Assemblage B specific Genus-specific

Unknown

Specificity

PCR, sequencing. This set of primer could combine with GDH-F3 and GDH-R3 in nested PCR PCR, RFLP, sequencing

Nested PCR, sequencing

Nested PCR, sequencing

Nested PCR, RFLP, sequencing

PCR, sequencing

Nested PCR, sequencing

PCR

PCR

Nested PCR, RFLP, sequencing

Assay Type

Genotyping and subtyping

Genotyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping and subtyping Genotyping and subtyping

Genotyping and subtyping

Genotyping

Genotyping

Genotyping

Usage

Table 50.1 List of the Gene, Primer, Assay Type and Main use of some Published G. duodenalis Genotyping Tools

Human, animal (guinea pig, slow loris, rat)

Dog

Human, animal (cat, dog, pig, rat)

Human

Human

Human, animal (dog, cattle, muskrat, beaver, rat, rabbit) Human, dog

Human

Human

Human, animal (beaver, cat, guinea pig, dog)

Sample Type in Original Study

dog, cat, calf, monkey

Wastewater

Wastewater

Wastewater, Cattle, lamb

Sample Type in Other Studies

(Continued)

70

99,100

98

76,81

76,81

97

12,85,95,96

94

94

69

Reference

Giardia 705

GDH-F3: ATCTTCGAGAGGATGCTTGAG GDH-R3: AGTACGCGACGCTGGGATACT

GDH-F3: ATCTTCGAGAGGATGCTTGAG GDH-R4: TGTCCTTGCACATCTCCTCCA

GDH-F4: CGAGCGCGAGCCGAAGTATATCC GDH-R5: GATGTTYGCRCCCATCTGRTAGTTC

GDH-F5: TCAACGTYAAYCGYGGYTTCCGT GDH-R6: GTTRTCCTTGCACATCTCC GDH-F6: CAGTACAACTCYGCTCTCGG GDH-R6: SSU-F1: TCCGGTCGATCCTGCCGGA SSU-R1:GCTCTCCGGAGTCGAAC

SSU-F2: CATCCGGTCGATCCTGCC SSU-R2: AGTCGAACCCTGATTCTCCGCCCAGG

SSU-F3: AAGTGTGGTGCAGACGGACTC SSU-R3: CTGCTGCCGTCCTTGGATGT

SSU-F4: GACGCTCTCCCCAAGGAC SSU-R4: CTGCGTCACGCTGCTCG

SSU-F5 : GCGCACCAGGAATGTCTTGT SSU-R5 : TCACCTACGGATACCTTGTT

SSU-F6: TCCGGTYGATTCTGCC SSU-R6: CTGGAATTACCGCGGCTGCT

GDH

GDH

GDH

SSU rRNA

SSU rRNA

SSU rRNA

SSU rRNA

SSU rRNA

SSU rRNA

Primer (5′–3′)

GDH

Gene

Table 50.1  (Continued)

480

183

130

497

292

Unknown

Amplify most protozoan parasites

Unknown

Unknown

Genus-specific

Genus-specific

Genus-specific

432

298

Genus-specific

Genus-specific

Genus-specific

Specificity

1190

528

778

Size (bp)

PCR, sequencing

Nested PCR (combine with primers SSU-F2 and SSU-R2), sequencing Nested PCR (combine with primers SSU-F2 and SSU-R2), sequencing PCR, sequencing

PCR, sequencing

PCR, RFLP, sequencing

Semi-nested PCR, RFLP

PCR, sequencing

PCR, RFLP, sequencing

PCR, RFLP sequencing

Assay Type

Genotyping and subtyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping

Genotyping

Genotyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping and subtyping

Usage

Human, animal (beaver, cat, guinea pig, muskrat, dog, sheep, mouse, blue heron) Human, animal (cat, dog, pig, rat)

Human, dog

Cattle

Human, dog

Human, wastewater, animal (dog, cat, deer mouse)

Human, animal (cat, dog, cattle)

Human, animal (dog, cat, cattle)

Animal (dog, sheep, goat, roe deer)

Human

Sample Type in Original Study

Cat

Ferret, cattle, cat

Wastewater

Ferret, cattle

Sample Type in Other Studies

98

66

97,106

109

14,101, 106–108

68,77

11,81

105

104

101–103 

Reference

706 Molecular Detection of Foodborne Pathogens

EF1-α-F1: GCCGAGGAGTTCGACTACATC EF1-α-R1: GACGCCSGAGATCTTGTAGAC

EF1-α-F2: GCTCSTTCAAGTACGCGTGG EF1-α-R2: GCATCTCGACGGATTCSACC VSP-F1: GTTGGATCCGCCACCCAGCAGGTCGGCGGCCTAATG VSP-R1: GTATCC(T/C)GGAGCTCAC(G/A)CCTTACATGTTGTAGCTGCTCCA VSP-F2: CGTATCGATGAGCTGAGAAGGATTTTAATG VSP-R1: GTATCC(T/C)GGAGCTCAC(G/A)CCTTACATGTTGTAGCTGCTCCA VSP-F3: ACGACGGTACTAAGGGCACGTG VSP-R1: GTATCC(T/C)GGAGCTCAC(G/A)CCTTACATGTTGTAGCTGCTCCA VSP-F4: TTACTCTAGAATAAGCTGAGGGAAA-CTTCAATG VSP-R2: ACGCCTCTAGACYRCANABGAACCA-CCARCA where B = T, C or G, R = A or G, Y = C or T, and N = A, T, C or G

Ef1-α

Ef1-α

VSP

VSP

VSP

VSP

β-Giardin

β-Giardin

β-Giardin

Giardin-F1: AAGTGCGTCAACGAGCAGCT Giardin-R1: TTAGTGCTTTGTGACCATCGA Giardin-F2: CATAACGACGCCATCGCGGCTCTCAGGAA Giardin-R2: TTTGTGAGCGCTTCTGTCGTGGCAGCGCTA Giardin-F3: AAGCCCGACGACCTCACCCGCAGTGC Giardin-R3: GAGGCCGCCCTGGATCTTCGAGACGAC Giardin-F4: CATAACGACGCCATCGCGGCTCTCAGGAA Giardin-R3: Giardin-F5: GAACGAACGAGATCGAGGTCCG Giardin-R4: CTCGACGAGCTTCGTGTT

β-Giardin

Unknown

650

1619

520

1755

Gene vsp 1267-specific

Gene tsa 417 and tsp 411-specific

Gene tsp 11-specific

Gene tsa 417-specific

Unknown

191

1908

Unknown

Unknown

G. duodenalisspecific

Genus-specific

511

384

753

218

171

PCR, RFLP, sequencing

PCR, RFLP, sequencing

PCR, RFLP sequencing

PCR, RFLP, sequencing

PCR, sequencing

Nested PCR (combine with Giardin-F3 and Giardin-R3), RFLP sequencing PCR, sequencing

Semi-nested PCR, RFLP, sequencing

PCR, hybridization

PCR, hybridization

Genotyping and subtyping

Genotyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping and subtyping

Genotyping

Genotyping

Human, animal (sheep, dog, calves)

Human

Human, animal (sheep, dog, calves)

Human, animal (sheep, dog, calves)

Human, animal (cat, dog, pig, rat)

Human, dog

Human, animal (dog, cat, calf)

Human

Cattle, human

Cattle, human

Animal (dog, cat), wastewater

Water

(Continued)

113

67

113

113

98

97

112

71,81,111

110

83,110

Giardia 707

VSP-F5: TTGCTCGAGCAACTGGAAATGAA-GGTGGGTG VSP-R2: ACGCCTCTAGACYRCANABGAACCA-CCARCA where B = T, C or G, R = A or G, Y = C or T, and N = A, T, C, or G rDNA-F1: CGATAGCAGGTCTGTGATGC rDNA-R1: GCTACTGATATGCTTAAGTTC A-F1: CTGCAGGGGCAAGGCGTAGAT A-R1: CCACCGTGCCAGTCTTCTGGG B-F1: CTGCAGTAACACTGGCAAG B-F2: CTGCAGAGTTCTCCGCAGCG

VSP

1500

800

615

1290

Size (bp)

Assemblage A-specific Assemblage B-specific

Genus-specific

Gene vsp 1267-specific

Specificity

PCR, RFLP, sequencing PCR, RFLP sequencing

PCR, sequencing

PCR, RFLP sequencing

Assay Type

Subtyping

Subtyping

Genotyping

Genotyping and subtyping

Usage

Human

Human

Cattle

Human, animal (sheep, dog, calves)

Sample Type in Original Study

Animal

Animal

Sample Type in Other Studies

Notes: TPI, triose phosphate isomerase; RFLP, restriction fragment length polymorphism; GDH, glutamate dehydrogenase; EF-1α, elongation factor 1α; VSP, variant surface protein.

Unknown

Unknown

rRNA

Primer (5′–3′)

Gene

Table 50.1  (Continued)

102,104

102,104

114

113

Reference

708 Molecular Detection of Foodborne Pathogens

Giardia

variable 5′ and 3′ ends of the SSU rRNA locus are targeted for genotyping the relatively closely related assemblages, whereas the more conserved regions would only provide sufficient information for the differentiation of Giardia species but not for G. duodenalis assemblages. Using the current consensus sequence information, different genes and gene regions can be targeted for different genotyping applications. Older primers can be reassessed for their suitability for their original task, but changing the location of the primers can potentially change their scope and/or specificity. For example, the use of primers of Hopkins et al.14 for a 292-bp product on cat isolates may not be appropriate (as previously mentioned) due to their limited differentiation resolution. Likewise, the use of the GDH primer (GDH-F5 in Table 50.1) described for the discrimination of all genotypes11 may no longer be true, as two additional substitutions in assemblages D, E, and F have been discovered, potentially altering the primer’s differentiation power. Subtyping tools. Subgrouping within the zoonotic assemblages A and B is of interest as there may be a link between specific subgroups and their hosts, with particular attention to those that may infect humans. It is therefore important to establish the specific subgroups, based on their intra-genotypic substitution patterns, and the continuity of these subgroups across genetic loci. The AI and AII subgroups are quite robust.13 In the SSU rRNA gene, they are differentiated at the 3′ end by a single substitution.66 At the GDH locus, there are numerous AI/AII specific polymorphisms and the majority of isolates can be easily differentiated.13 The TPI gene has only two AI/AII specific polymorphisms in the region examined and there are no outliers,13 but has other nucleotide substitutions defining other groups. It is noteworthy that all these loci have produced consistent distinction between the two close subgroups. The conserved β-giardin alignment in contrast had only one substitution clearly differentiating the AI/AII subgroups and several other substitutions producing additional groups.13 The significance of the additional TPI and β-giardin subgroup within the assemblage A and the intergenic linkage of them are not yet clear. In fact, these two genes have consistently demonstrated numerous subgroups within all assemblages. Their significance, however, is difficult to determine because the samples that had been characterized at other loci did not always segregate into the same subgroups defined by the β-giardin locus. The BIII and BIV subgroups originally described by allozyme electrophoretic studies80 are not reproducible in DNA sequence analyses.13 At the SSU rDNA locus, there are insufficient full-length samples characterized previously as BIII or BIV to assess if there are in fact any BIII/BIV specific polymorphisms present. At the β-giardin locus, there are also insufficient reference isolates previously characterized as BIII/BIV to determine which substitutions may be BIII/BIV specific. Attempts to use isolates characterized at other loci (and β-giardin) as references have led to conflicting results, as samples are usually grouped differently at the different loci.81 At the GDH and TPI loci, the BIII and BIV isolates appeared to form the most distant subgroups within

709

assemblage B. However, this subgrouping of assemblage B is clouded with extra substitutions forming additional groupings, which also share some of the substitutions found in BIII and BIV. Thus, in assemblage B, there are more subgroups than just BIII and BIV. It is also interesting to note, that with the increasing number of samples characterized, subgroups have been identified in assemblage E. Assemblage E, like assemblages A and B, has a broad host range (various hoofed livestock), therefore, may contain host-specific subgroups. The subgrouping is again most obvious at the β-giardin locus. The nucleotide sequences do not yet appear to segregate in the host-specific manner as the allozyme data have demonstrated,9 but the number of representative sequences from the different livestock hosts is still small. It is not possible to speculate on subgroupng in the other assemblages (C, D, F, and G) as there are too few data available. For assemblage D, there is one apparently host-specific polymorphism in the β-giardin alignment where samples have included dogs and a coyote described by Santin et al.82 In contrast, there is no host-specific polymorphism in assemblage G, including rats and a mouse in the GDH alignment.13 Molecular tools for the analysis of environmental samples. Diagnosis of Giardia to the species/genotype level is especially a challenge for environmental samples because of the usual low number of cysts and the rich presence of PCR inhibitors. The performance of some PCR methods in the analysis of environmental samples has been evaluated with Giardia negative samples seeded with known numbers of G. muris cysts. PCR was performed on DNA extracted directly from water concentrates seeded with Giardia cysts. The inhibitory effect of surface water on PCR was reported.83 In one study, the inhibition effect was removed by purifying the extracted DNA from wastewater by FastDNA kit (Q-Biogene, Carlsbad, CA).84 More recent techniques have used a cyst isolation step using IMS prior to DNA extraction to remove PCR inhibitors or contaminants present in water samples.83,85 Thus, the molecular methods used for studying Giardia species or G. duodenalis assemblages that are present in the environment are mainly adapted from methods used for fecal samples following an IMS procedure. A strategy combining direct DNA extraction using the FastDNA soil DNA extraction kit and neutralization of residual PCR inhibitors using high concentrations of nonacetylated bovine serum albumin in PCR has been used successfully in the PCR detection of Cryptosporidium oocysts in water samples86 and this DNA extraction method has been used effectively in our laboratories in PCR detection of Giardia cysts in water samples. There are only five reports available on molecular identification of Giardia in wastewater samples. The PCR-RFLP or sequence analyses of the β-giardin, TPI, SSU rRNA and GDH genes were used in the molecular detection in wastewater concentrates from Italy,84 USA,85 Canada,77 Norway,87 and Mexico.88 In all wastewater samples examined, only G. duodenalis cysts were identified and, more importantly, only Giardia pathogens of humans (assemblages A and B) were

710

Molecular Detection of Foodborne Pathogens

found. To date, no study has been performed on molecular detection of Giardia in food.

50.2  Methods 50.2.1 Reagents and Equipment 50.2.1.1 Supplies for Sample Collection and Preservation (1) 10-L or 100-L carboy (2) Centrifuge tubes: conical, graduated, 1.5, 15-ml (3) Potassium dichromate 50.2.1.2 Supplies for Filtration and Centrifugation (1) Stomacher bag, Seward, London, UK (2) Envirochek capsule filters (product no. 12110), Pall Gelman Laboratory, MI (3) Envirochek HV capsule filter (product no. 12099), Pall Gelman Laboratory (4) Laboratory shaker with arms (5) Centrifuge: capable of accepting 250-ml conical centrifuge tubes and achieving 1,500 × g (6) Centrifuge tubes: conical, graduated, 50, and 250-ml (7) Chemicals: glycine, Laureth-12, Tris, EDTA, antifoam A 50.2.1.3 Supplies for IMS (1) Dynabeads GC-Combo kit (product No. 730.12), Dynal A.S, Oslo, Norway (2) Dynal Magnetic Particle Concentrators (Dynal MPC): Dynal MPC-S (product no. 120.20) and Dynal MPC-1 (product no. 120.01), Dynal A.S, Oslo, Norway (3) Flat-sided sample tube (product no. 740.03), Dynal L10, Dynal A.S, Oslo, Norway 50.2.1.4 Supplies for DNA Extraction (1) QIAamp DNA mini kit: product no. 51304 (50 tests) or 51306 (250 tests), Qiagen Inc., Valencia, CA (2) QIAamp DNA Stool mini kit: product no. 51504 (50 tests) or 51306 (250 tests), Qiagen Inc., Valencia, CA (3) FastDNA SPIN kit (for soil), Q-BIOgene Corporation (1-760-929-1700). Catalog number 6560-200. Size: 50 reactions (4) Equipment: FastPrep® Instrument, Q-BIOgene (5) Chemicals: KOH, dithiothreitol, phenol, chloroform, isoamyl alcohol 50.2.1.5 Supplies for PCR (1) Primary TPI PCR primers: (a) Forward (TPI-F5 in Table 5′-AAATIATGCCTGCTCGTCG-3′ (b) Reverse (TPI-R5 in Table 5′-CAAACCTTITCCGCAAACC-3′

50.1): 50.1):

(2) Secondary TPI PCR primers: (a) Forward (TPI-F6 in Table 5′-CCCTTCATCGGIGGTAACTT-3′ (b) Reverse (TPI-R6 in Table 5′-GTGGCCACCACICCCGTGCC-3′ (3) 10 × PCR buffer (4) 100 mM dNTP (5) Taq polymerase (6) Albumin, bovine serum, nonacetylated (7) 25 mM MgCl2 (8) 100 bp DNA ladder

50.1): 50.1):

50.2.1.6 Supplies for DNA Sequencing (1) 3130 AB Prism Genetic Analyzer (2) BigDye® Terminator V3.1 Cycle Sequencing Kit (3) 96-well reaction plate and septa

50.2.2 Sample Preparation The following Giardia genotyping and subtyping techniques are used in the Molecular Epidemiology Laboratory at the Division of Parasitic Diseases, Centers for Disease Control and Prevention. Giardia cysts in 30 g fresh vegetable or salad are processed through stomaching following Cook et al.65 Giardia cysts in 10-L of surface water or 100-L finished water samples are concentrated through filtration and elution following USEPA method 1623.64 The centrifugation and IMS steps for both food and water samples are processed following USEPA method 1623.64 For fecal specimens, DNA is extracted using either the QIAamp® DNA Stool Kit (QIAGEN Inc, Valencia, CA)89 or the FastDNA® SPIN Kit for Soil Samples (Q-BIOgene, Irvine CA).86 The latter requires the use of a FastPrep instrument, which may not be available in some laboratories. For food and water samples, DNA is extracted using QIAamp DNA mini kit (Qiagen) after the cysts are isolated by IMS using Dynabeads® GC combo kit (Dynal, Oslo, Norway)90 or directly without cyst isolation using the FastDNA® SPIN Kit for Soil Samples, with the latter technique slightly less sensitive but more economic.86 50.2.2.1  Fecal Sample Collection and Preservation Store at least 2 g of fecal specimens containing Giardia cysts unpreserved at 4°C, or in 2.5% potassium dichromate solution at 4°C, or frozen at –20°C. Proceed directly to DNA extraction as specified in Section 50.2.2.5. 50.2.2.2 Extraction of Giardia Cysts from Food and Water Samples (i) Food sample: (1) Collect and process 30 g of fresh vegetables or salad as soon as possible. (2) Place the food in a filtered stomacher bag (Seward, London, UK), add 200 ml of 1 M glycine pH 5.5 to the bag, and stomach the sample for 30 sec to elute cysts from fresh vegetable surfaces.

Giardia

(3) Pull the filter upward in the stomach bag to remove the sample from the extractant and squeeze by hand to remove as much of the extractant as possible. Discard the filter bag containing the sample. (ii) Water sample: (1) Collect and process 10 L of surface water samples or 100 L of finished water samples as soon as possible. (2) Filter the water samples at a flow rate of 2.0 L/min through Envirochek capsule filters or Envirochek HV capsule filters (Pall Gelman Sciences). The Envirochek HV filter is preferred for finished water samples. (3) Elute cysts from the capsule filters with elution buffer and wrist action agitation, as specified in EPA Method 1623. 50.2.2.3 Centrifugation of Giardia Cysts from Food and Water Samples (1) Collect the extractant or eluant in 250-ml conicalbottom centrifuge tubes, and concentrate the cysts by centrifugation at 1,500 × g for 15 min. (2) Record the resulting pellet volumes and process 0.5 ml or less of the pellet with DNA extraction using the FastDNA SPIN Soil kit as specified below. If the QIAamp DNA mini kit is used in DNA extraction, cyst isolation using the IMS protocol below is needed prior to the DNA extraction. 50.2.2.4 Purification of Giardia Cysts from Food and Water Samples (1) Wash the packed pellet (0.5 ml of packed pellet volume or less) in 15 ml polypropylene tube twice with distilled water by centrifugation at 1,500 × g for 10 min. (2) Isolate Giardia cysts by IMS using the Dynabeads® GC combo kit (Dynal A.S, Oslo, Norway) and manufacturer-suggested procedures. 50.2.2.5 DNA Extraction from Fecal Specimens, Food, and Water Samples (i) Extraction of DNA in Fecal Specimens using the QIAamp DNA Stool Kit: (1) Transfer ~200 µl of fecal specimen to a 2.0-ml microfuge tube. (2) Wash at least twice with distilled water by centrifugation at 1,000 × g for 10 min. By-pass the washing step if the fecal specimen is stored frozen. (3) Add 66.6 µl of 1 M KOH, and 18.6 µl of 1 M dithiothreitol (DTT) to the pellet, and mix thoroughly by stirring with a pipette tip. (4) Incubate at 65°C for 15 min. (5) Neutralize the alkaline with 8.6 µl of 25% HCl and 160 µl of 2 M Tris-HCl (pH 8.3). Vortex the tube.

711

(6) Add 250 µl of phenol:chloroform:isoamyl alcohol (25:24:1) solution to the tube and vortex. (7) Centrifuge at 500 × g for 5 min. (8) Transfer the supernatant to a 2.0 ml microfuge tube. (9) Add 1 ml of ASL buffer from the QIAamp® DNA Stool Kit (QIAGEN Inc, Valencia, CA) to the supernatant. Incubate the mixture at 80°C for 5 min. (10) Add 1 InhibitEX tablet to each specimen and vortex immediately for 1 min or until the tablet is completely dissolved. (11) Follow the remaining procedures specified in the QIAamp® DNA Stool Kit. Elute the DNA using 200 µl of the AE Elution Buffer, and store the extractant at –20°C. (ii) Extraction of Giardia DNA from IMS Purified Cysts from Water and Food Samples using the QIAamp® DNA Mini Kit: (1) Add 180 µl of Buffer ATL from the QIAamp DNA Mini Kit (Qiagen) to the 2.0 ml microfuge tube containing magnetic beads-Giardia cysts (without the cyst detachment process), and vortex for 30 sec. (2) Freeze-thaw five times at –70°C (or dry ice) and 56°C. (3) Add 20 µl of proteinase K from the kit to the tube, vortex for 10 sec, and incubate at 56°C overnight. (4) Add 200 µl of buffer AL to the tube, vortex, and incubate the tube at 70°C for 10 min. (5) Follow the remaining procedures specified in the QIAamp DNA Mini Kit. Elute the DNA using 100 µl of buffer AE, and store the extractant at –20°C. (iii) Extraction of DNA from Fecal Specimen, Food and Water Concentrates using the FastDNA® SPIN Soil Kit: (1) Wash 0.5 ml of packed food and water concentrates or fecal suspension in 2.0 ml microfuge tubes twice with distilled water by centrifugation at 10,000 × g for 3 min. (2) Aspirate the supernatant. Use the pellet immediately for DNA extraction. (3) Re-suspend the washed water or fecal pellet in 978 µl of sodium phosphate buffer from the FastDNA SPIN Soil Kit (Q-BIOgene, Irvine, CA). (4) Transfer the suspension to a lysing matrix E tube. (5) Add 122 µl of MT buffer to the lysing matrix E tube. (6) Secure the tube in FastPrep® Instrument (Q-BIOgene) and process for 30 sec at the speed setting 5.5. (7) Centrifuge the lysing matrix E tube at 10,000 × g for 30 sec. (8) Transfer supernatant to a clean tube. Add 250 µl of PPS reagent from the kit and mix by shaking the tube by hand ten times.

712

Molecular Detection of Foodborne Pathogens

(9) Follow the remaining procedures specified in the FastDNA® SPIN Soil Kit. Elute the DNA using 100 µl of DES (DNase/pyrogen free water), and store the extraction at –20°C.

50.2.3  Detection Procedures Giardia cysts are first detected by nested PCR analysis of the TPI gene. G. duodenalis genotypes and subtypes are determined by sequence analysis of the secondary PCR products. Fecal specimens are generally analyzed twice with 2 µl of extracted DNA, whereas food and water samples are each analyzed five times with 2 µl of extracted DNA because of the low number of cysts and likely presence of mixed genotypes. Positive (DNA of genotypes or subtypes unexpected in specimens under analysis) and negative (reagent water) are used in each PCR run. In the protocol described below, the primers TPI-F5, R5, F6, and R6 in Table 50.1 are used in nested PCR analysis of the Giardia TPI gene. TPI-F5 and R5 are used in primary PCR whereas TPI-F6 and R6 are used in the secondary PCR. 50.2.3.1  Primary PCR (1) For each PCR reaction, prepare the following ­master mixture: 10 × Perkin-Elmer PCR buffer dNTP (1.25 mM)

10 µl

TPI-F5 primer (40 ng/µl)

2.5 µl

TPI-R5 primer (40 ng/µl) MgCl2 (25 mM)

6 µl

16 µl 2.5 µl

Nonacetylated bovine serum albumin (10 mg/ml) Distilled water

4 µl

Taq polymerase

0.5 µl

Total

98 µl

56.5 µl

(2) Add 98 µl of the master mixture to each PCR tube. (3) Add 2 µl of DNA suspension to each tube. (4) Run the following PCR program: one cycle of 94°C for 3 min; 35 cycles of: 94°C for 45 sec, 50°C for 45 sec and 72°C for 1 min; and one cycle of 72°C for 7 min followed by 4°C soaking. 50.2.3.2  Secondary PCR (1) For each PCR reaction, prepare the following master mixture: 10 × Perkin-Elmer PCR buffer dNTP (1.25 mM)

10 µl

TPI-F6 primer (40 ng/µl)

5 µl

TPI-R6 primer (40 ng/µl) MgCl2 (25 mM)

6 µl

Distilled water

55.5 µl

Taq polymerase

0.5 µl

Total

98 µl

16 µl 5 µl

(2) Add 98 µl of the master mixture to each PCR tube. (3) Add 2 µl of the primary PCR reaction to each tube. (4) Run the following PCR program: one cycle of 94°C for 3 min; 35 cycles of: 94°C for 45 sec, 50°C for 45 sec and 72°C for 1 min; and one cycle of 72°C for 7 min followed by 4°C soaking. 50.2.3.3 Detection of Secondary PCR Products Run electrophoresis on 1.5% agarose gel with 15 µl of the secondary PCR product, using the 100-bp ladder as the control. 50.2.3.4 DNA Sequencing (1) Clean the positive secondary PCR products of the expected size using the Montage-PCR kit (Millipore, Bedford, MA). The cleaned PCR products can be stored at –20°C. (2) Sequence the cleaned PCR products using primers TPI-F6 and TPI-R6. (3) For each sequencing reaction, prepare the following using components of the BigDye® Terminator V3.1 Cycle Sequencing Kit (Applied Biosystems, Foster, CA): BigDye terminator buffer

4 µl

BigDye terminator

1 µl

Primer (40 ng/µl) Reagent water

2 µl 9.5 µl

(4) Add 19.5 µl of the master mixture to a PCR tube. (5) Add 0.5 µl cleaned PCR products to each tube. (6) Run the following program in a PCR machine: 25 cycles of 96°C for 10 sec, 50°C for 5 sec, and 60°C for 4 min followed by 4°C soaking. (7) Clean the 20 µl of sequencing reaction products using the Centrisep Spin Column (Princeton Separations, Adelphia, NJ). (8) Dry the cleaned sequencing products in a vacuum centrifuge. The cleaned products can be stored in a freezer. (9) Add 15 µl of BigDye Formamide to the dehydrated sequencing products. (10) Transfer the DNA suspension to the MicroAmp Optical 96-well reaction plate (Applied Biosystems). (11) Cover the plate with a 96-well Plate Septa (Applied Biosystems). (12) Heat the plate in a PCR machine at 94°C for 5 min, and cool the plate at –20°C for at least 3 min. (13) Load the reaction product onto a 3130 AB Prism Genetic Analyzer for sequencing using proper program. 50.2.3.5 Sequence Analysis (1) Read out the electropherograms generated by the sequencer using the ChromasPro software (www. technelysium.com.au/ChromasPro.html) or any other software.

713

Giardia

(2) Align the TPI sequences generated with each other and reference sequences using the software ClustalX (ftp://ftp-igbmc.u-strasbg.fr/pub/ClustalX/). (3) Check the sequence alignment for sequencing accuracy using the software BioEdit (www.mbio.ncsu. edu/BioEdit/bioedit.html). (4) Recheck the electropherograms for any sequence uncertainty. (5) Determine G. duodenalis genotypes and subtypes based on sequence identity to reference sequences.

50.3 Conclusions and Further Perspective Molecular tools have been used to provide clearer insights into the taxonomy, host range and transmission routes of Giardia and the epidemiology of human giardiasis.91 These tools are essential for the characterization of the parasite at species, assemblage, and subtype levels, and the assessment of the public health importance of different routes of transmission. Specific aspects of Giardia biology—including their low infectious doses, host specificity and host adaptation, and the prolonged survival of cysts in moist environments—have particular relevance in determining transmission routes. To maximize the use of molecular epidemiological tools, they must be applicable to both clinical (human and nonhuman) and environmental (including food) samples, preferably from the same community or area, for species and genotype identification, transmission dynamics characterization, human infection potential assessment, and infection/contamination source tracking. An ongoing issue in Giardia research is the question of zoonotic transmission.7,92,93 Data accumulated so far have shown only limited potential of zoonotic transmission of giardiasis, especially infections caused by assemblage A. Nevertheless, the number of human and animal studies that have used genotyping and subtyping tools is still small, which does not allow the generation of firm conclusions. This is especially true for assemblage B, whose prevalence in animals is not well known, and population structure is not clear. Clearly, infections with assemblage B are common in some animals that have long been considered sources of human infections, such as beavers and muskrats.12 Whether there is any host adaptation within the assemblage B remains to be answered. Likewise, answers to questions on the significance of other subtypes identified at the TPI and β-giardin loci within the assemblage A and their linkage across genetic loci will have important implications in assessing disease burdens attributable to zoonotic transmission. The application of molecular biologic tools in the analysis of Giardia cysts in environmental samples is still in its infancy and has really not demonstrated potential as in the case of the sister parasite Cryptosporidium. Thus far, there are only a handful of studies that have genotyped Giardia cysts in water samples and there is no study on food, juice, or soil samples. Results of the studies of water samples have so far shown the presence of only G. duodenalis assemblages

A and B. This conclusion likely will change in the future, as other Giardia species and G. duodenalis assemblages will most probably be found in future studies, especially G. duodenalis assemblage E in surface or source water. Identifying contamination sources of Giardia cysts in the environment requires an in-depth understanding of host specificity of the parasite at both the genotype and subtype levels, which reinforces the need for more extensive characterizations of Giardia in various animals. Because the numbers of cysts recovered from water and food matrices are likely to be small, the sensitivity of current PCR tools has to be improved, and new strategies need to be developed to maximize the efficiency of DNA extraction and minimize the co-extraction and effects of PCR inhibitors.

Acknowledgments Y. Feng acknowledges the financial support from the National Natural Science Foundation of China (No. 30771881). The findings and conclusions in this report are those of the authors and do not necessarily represent the views of the Centers for Disease Control and Prevention.

References



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714 15. Monis, P.T. and Andrews, R.H. Molecular epidemiology: assumptions and limitations of commonly applied methods. Int. J. Parasitol., 28, 981, 1998. 16. Leonhard, S. et al. The molecular characterisation of Giardia from dogs in southern Germany. Vet. Parasitol., 150, 33, 2007. 17. Ey, P.L. et al. Genetic analysis of Giardia from hoofed farm animals reveals artiodactyl-specific and potentially zoonotic genotypes. J. Eukaryot. Microbiol., 44, 626, 1997. 18. Smith, H.V. et al. Cryptosporidium and Giardia as foodborne zoonoses. Vet. Parasitol., 149, 29, 2007. 19. Karanis, P., Kourenti, C. and Smith, H. Waterborne transmission of protozoan parasites: a worldwide review of outbreaks and lessons learnt. J. Water Health, 5, 1, 2007. 20. Savioli, L., Smith, H. and Thompson, A. Giardia and Cryptosporidium join the ‘Neglected Diseases Initiative’. Trends Parasitol., 22, 203, 2006. 21. Amahmid, O., Asmama, S. and Bouhoum, K. The effect of waste water reuse in irrigation on the contamination level of food crops by Giardia cysts and Ascaris eggs. Int. J. Food Microbiol., 49, 19, 1999. 22. Robertson, L.J. and Gjerde, B., Isolation and enumeration of Giardia cysts, Cryptosporidium oocysts, and Ascaris eggs from fruits and vegetables. J. Food Prot., 63, 775, 2000. 23. Robertson, L.J. and Gjerde, B. Occurrence of Cryptosporidium oocysts and Giardia cysts in raw waters in Norway. Scand. J. Public Health, 29, 200, 2001. 24. Robertson, L.J. et al. Microbiological analysis of seed sprouts in Norway. Int. J. Food Microbiol., 75, 119, 2002. 25. Chaidez, C. et al. Occurrence of Cryptosporidium and Giardia in irrigation water and its impact on the fresh produce industry. Int. J. Environ. Health Res., 15, 339, 2005. 26. Thurston-Enriquez, J.A. et al. Detection of protozoan parasites and microsporidia in irrigation waters used for crop production. J. Food Prot., 65, 378, 2002. 27. Robertson, L.J. and Gjerde, B. Factors affecting recovery efficiency in isolation of Cryptosporidium oocysts and Giardia cysts from vegetables for standard method development. J. Food Prot., 64, 1799, 2001. 28. Robertson, L.J. and Gjerde, B. Occurrence of parasites on fruits and vegetables in Norway. J. Food Prot., 64, 1793, 2001. 29. Osterholm, M.T. et al. An outbreak of foodborne giardiasis. N. Engl. J. Med., 304, 24, 1981. 30. Petersen, L.R., Cartter, M.L. and Hadler, J.L. A food-borne outbreak of Giardia lamblia. J. Infect. Dis., 157, 846, 1988. 31. Quick, R. et al. Restaurant-associated outbreak of giardiasis. J. Infect. Dis., 166, 673, 1992. 32. Conroy, D. A note on the occurrence of Giardia sp. in a Christmas pudding. Rev. Iber. Parasitol. 20, 567, 1960. 33. Karabiber, N. and Aktas, F. Foodborne giardiasis. Lancet, 337, 376, 1991. 34. Hlavsa, M.C., Watson, J.C. and Beach, M.J. Giardiasis surveillance—United States, 1998–2002. MMWR Surveill. Summ., 54(1), 9, 2005. 35. Yoder, J.S. and Beach, M.J. Giardiasis surveillance—United States, 2003–2005 MMWR Surveill. Summ., 56(7), 11, 2007. 36. Huang, D.B. and White, A.C. An updated review on Cryptosporidium and Giardia. Gastroenterol. Clin. North Am., 35, 291, 2006. 37. Caeiro, J.P. et al. Etiology of outpatient pediatric nondysenteric diarrhea: a multicenter study in the United States. Pediatr. Infect. Dis. J., 18, 94, 1999. 38. Gilman, R.H. et al. Epidemiology and serology of Giardia lamblia in a developing country: Bangladesh. Trans. R. Soc. Trop. Med. Hyg., 79, 469, 1985.

Molecular Detection of Foodborne Pathogens 39. Fraser, D. et al. Natural history of Giardia lamblia and Cryptosporidium infections in a cohort of Israeli Bedouin infants: a study of a population in transition. Am. J. Trop. Med. Hyg., 57, 544, 1997. 40. Farthing, M.J. et al. Natural history of Giardia infection of infants and children in rural Guatemala and its impact on physical growth. Am. J. Clin. Nutr., 43, 395, 1986. 41. Gilman, R.H. et al. Rapid reinfection by Giardia lamblia after treatment in a hyperendemic Third World community. Lancet, 1, 343, 1988. 42. Cifuentes, E. et al. Risk factors for Giardia intestinalis infection in agricultural villages practicing wastewater irrigation in Mexico. Am. J. Trop. Med. Hyg., 62, 388, 2000. 43. Lee, S.H. et al. Surveillance for waterborne-disease ­outbreaks—United States. 1999–2000. MMWR Surveill. Summ., 51, 1, 2002. 44. Stuart, J.M. et al. Risk factors for sporadic giardiasis: a casecontrol study in southwestern England. Emerg. Infect. Dis., 9, 229, 2003. 45. Thompson, S.C. Giardia lamblia in children and the child care setting: a review of the literature. J. Paediatr. Child Health, 30, 202, 1994. 46. Hill, D.R. Giardiasis. Issues in diagnosis and management. Infect. Dis. Clin. North Am., 7, 503, 1993. 47. Olsen, S.J. et al. Surveillance for foodborne-disease ­outbreaks—United States, 1993–1997. MMWR CDC Surveill. Summ., 49, 1, 2000. 48. Ekdahl, K. and Andersson, Y. Imported giardiasis: impact of international travel, immigration, and adoption. Am. J. Trop. Med. Hyg., 72, 825, 2005. 49. Xiao, L. and Fayer, R. Molecular characterization of species and genotypes of Cryptosporidium and Giardia and assessment of zoonotic transmission. Int. J. Parasitol., 38, 1239, 2008. 50. Chin, A.C. et al. Strain-dependent induction of enterocyte apoptosis by Giardia lamblia disrupts epithelial barrier function in a caspase-3-dependent manner. Infect. Immun., 70, 3673, 2002. 51. Scott, K.G. et al. Intestinal infection with Giardia spp. reduces epithelial barrier function in a myosin light chain kinase-dependent fashion. Gastroenterology, 123, 1179, 2002. 52. Buret, A.G. et al. Giardia lamblia disrupts tight junctional ZO-1 and increases permeability in non-transformed human small intestinal epithelial monolayers: effects of epidermal growth factor. Parasitology, 125, 11, 2002. 53. Nash, T.E. Surface antigenic variation in Giardia lamblia. Mol. Microbiol., 45, 585, 2002. 54. Touz, M.C. Conrad, J.T. and Nash, T.E., A novel palmitoyl acyl transferase controls surface protein palmitoylation and cytotoxicity in Giardia lamblia. Mol. Microbiol., 58, 999, 2005. 55. Hiatt, R.A., Markell, E.K. and Ng, E. How many stool examinations are necessary to detect pathogenic intestinal protozoa? Am. J. Trop. Med. Hyg., 53, 36, 1995. 56. Mank, T.G. et al., Sensitivity of microscopy versus enzyme immunoassay in the laboratory diagnosis of giardiasis. Eur. J. Clin. Microbiol. Infect. Dis., 16, 615, 1997. 57. Aldeen, W.E. et al. Comparison of nine commercially available enzyme-linked immunosorbent assays for detection of Giardia lamblia in fecal specimens. J. Clin. Microbiol., 36, 1338, 1998. 58. Garcia, L.S. and Shimizu, R.Y. Evaluation of nine immunoassay kits (enzyme immunoassay and direct fluorescence) for detection of Giardia lamblia and Cryptosporidium parvum in human fecal specimens. J. Clin. Microbiol., 35, 1526, 1997.

Giardia 59. Zimmerman, S.K. and Needham, C.A. Comparison of conventional stool concentration and preserved-smear methods with Merifluor Cryptosporidium/Giardia Direct Immunofluorescence Assay and ProSpecT Giardia EZ Microplate Assay for detection of Giardia lamblia. J. Clin. Microbiol., 33, 1942, 1995. 60. Chan, R. et al. Evaluation of a combination rapid immunoassay for detection of Giardia and Cryptosporidium antigens. J. Clin. Microbiol., 38, 393, 2000. 61. Aziz, H. et al. A comparison study of different methods used in the detection of Giardia lamblia. Clin. Lab. Sci., 14, 150, 2001. 62. Water Supply (Water Quality) (Amendment) Regulations 1999. SI No. 1524., 1999. 63. DWI Information Letter. The Water Supply (Water Quality) (Amendment) Regulations 2000, SI No. 3184 England 2001, SI No. 3911 (W.323) Wales: Cryptosporidium in Water Supplies, Laboratory and Analytical Procedures. Part 2, June 2005. UK Drinking Water Inspectorate, 2005. 64. USEPA, Method 1623. Giardia and Cryptosporidium in water by filtration/IMS/FA. United States Environmental Protection Agency, Office of Water, Washington, DC. EPA 821-R01-025, 2001. 65. Cook, N. et al. Development of a method for detection of Giardia duodenalis cysts on lettuce and for simultaneous analysis of salad products for the presence of Giardia cysts and Cryptosporidium oocysts. Appl. Environ. Microbiol., 73, 7388, 2007. 66. Weiss, J.B., van Keulen, H. and Nash, T.E. Classification of subgroups of Giardia lamblia based upon ribosomal RNA gene sequence using the polymerase chain reaction. Mol. Biochem. Parasitol., 54, 73, 1992. 67. Ey, P.L. et al. Giardia intestinalis: detection of major genotypes by restriction analysis of gene amplification products. Int. J. Parasitol., 23, 591, 1993. 68. van Keulen, H. et al. A three nucleotide signature sequence in small subunit rRNA divides human Giardia in two different genotypes, J. Eukaryot. Microbiol., 42, 392, 1995. 69. Baruch, A.C., Isaac-Renton, J. and Adam, R.D. The molecular epidemiology of Giardia lamblia: a sequence-based approach, J. Infect. Dis., 174, 233, 1996. 70. Monis, P.T. et al. Molecular genetic analysis of Giardia intestinalis isolates at the glutamate dehydrogenase locus. Parasitology, 112, 1, 1996. 71. Caccio, S.M., De Giacomo, M. and Pozio, E. Sequence analysis of the beta-giardin gene and development of a polymerase chain reaction-restriction fragment length polymorphism assay to genotype Giardia duodenalis cysts from human faecal samples. Int. J. Parasitol., 32, 1023, 2002. 72. Sogin, M.L. et al. Phylogenetic meaning of the kingdom concept: an unusual ribosomal RNA from Giardia lamblia. Science, 243, 75, 1989. 73. van Keulen, H. et al. Unique phylogenetic position of Diplomonadida based on the complete small subunit ribosomal RNA sequence of Giardia ardeae, G. muris, G. duodenalis and Hexamita sp. Faseb. J., 7, 223, 1993. 74. van Keulen, H. et al. Nucleotide sequence of the 5.8S and large subunit rRNA genes and the internal transcribed spacer and part of the external spacer from Giardia ardeae. Nucleic Acids Res., 19, 6050, 1991. 75. Lu, S.Q., Baruch, A.C. and Adam, R.D. Molecular comparison of Giardia lamblia isolates. Int. J. Parasitol., 28, 1341, 1998. 76. Amar, C.F. et al. Sensitive PCR-restriction fragment length polymorphism assay for detection and genotyping of Giardia duodenalis in human feces. J. Clin. Microbiol., 40, 446, 2002.

715 77. van Keulen, H. et al. Presence of human Giardia in domestic, farm and wild animals, and environmental samples suggests a zoonotic potential for giardiasis. Vet. Parasitol., 108, 97, 2002. 78. Berrilli, F. et al. Genotype characterisation of Giardia duodenalis isolates from domestic and farm animals by SSU-rRNA gene sequencing. Vet. Parasitol., 122, 193, 2004. 79. Fayer, R. et al. Detection of Cryptosporidium felis and Giardia duodenalis Assemblage F in a cat colony. Vet. Parasitol., 140, 44, 2006. 80. Andrews, R.H. et al. Giardia intestinalis: electrophoretic evidence for a species complex. Int. J. Parasitol., 19, 183, 1989. 81. Robertson, L.J. et al. Application of genotyping during an extensive outbreak of waterborne giardiasis in Bergen, Norway, during autumn and winter 2004. Appl. Environ. Microbiol., 72, 2212, 2006. 82. Santin, M. et al. First report of Giardia in coyotes (Canis latrans). J. Eukaryot. Microbiol., 50 (Suppl), 709, 2003. 83. Mahbubani, M.H. et al. Detection of Giardia in environmental waters by immuno-PCR amplification methods. Curr. Microbiol., 36, 107, 1998. 84. Caccio, S.M. et al. Giardia cysts in wastewater treatment plants in Italy. Appl. Environ. Microbiol., 69, 3393, 2003. 85. Sulaiman, I.M. et al. Distribution of Giardia duodenalis genotypes and subgenotypes in raw urban wastewater in Milwaukee, Wisconsin. Appl. Environ. Microbiol., 70, 3776, 2004. 86. Jiang, J. et al. Development of procedures for direct extraction of Cryptosporidium DNA from water concentrates and for relief of PCR inhibitors. Appl. Environ. Microbiol., 71, 1135, 2005. 87. Robertson, L.J., Hermansen, L. and Gjerde, B.K. Occurrence of Cryptosporidium oocysts and Giardia cysts in sewage in Norway. Appl. Environ. Microbiol., 72, 5297, 2006. 88. Di Giovanni, G.D. et al. Investigation of potential zooanthroponotic transmission of cryptosporidiosis and giardiasis through agricultural use of reclaimed wastewater. Int. J. Environ. Health Res., 16, 405, 2006. 89. Xiao, L. et al. Host adaptation and host-parasite co-evolution in Cryptosporidium: implications for taxonomy and public health. Int. J. Parasitol., 32, 1773, 2002. 90. Xiao, L., Lal, A.A. and Jiang, J. Detection and differentiation of Cryptosporidium oocysts in water by PCR-RFLP. Methods Mol. Biol., 268, 163, 2004. 91. Smith, H.V. et al. Tools for investigating the environmental transmission of Cryptosporidium and Giardia infections in humans. Trends Parasitol., 22, 160, 2006. 92. Thompson, R.C. and Monis, P.T. Variation in Giardia: implications for taxonomy and epidemiology. Adv. Parasitol., 58, 69, 2004. 93. Hunter, P.R. and Thompson, R.C. The zoonotic transmission of Giardia and Cryptosporidium. Int. J. Parasitol., 35, 1181, 2005. 94. Bertrand, I., Albertini, L. and Schwartzbrod, J. Comparison of two target genes for detection and genotyping of Giardia lamblia in human feces by PCR and PCR-restriction fragment length polymorphism. J. Clin. Microbiol., 43, 5940, 2005. 95. Aloisio, F. et al. Severe weight loss in lambs infected with Giardia duodenalis assemblage B. Vet. Parasitol., 142, 154, 2006. 96. Feng, Y. et al. High intragenotypic diversity of Giardia duodenalis in dairy cattle on three farms, Parasitol. Res., 103, 87, 2008. 97. Traub, R.J. et al. Epidemiological and molecular evidence supports the zoonotic transmission of Giardia among humans and dogs living in the same community. Parasitology, 128, 253, 2004.

716 98. Monis, P.T. The importance of systematics in parasitological research. Int. J. Parasitol., 29, 381, 1999. 99. Abe, N., Kimata, I. and Iseki, M. Identification of genotypes of Giardia intestinalis isolates from dogs in Japan by direct sequencing of the PCR amplified glutamate dehydrogenase gene. J. Vet. Med. Sci., 65, 29, 2003. 100. Itagaki, T. et al. Genotyping of Giardia intestinalis from domestic and wild animals in Japan using glutamete dehydrogenase gene sequencing. Vet. Parasitol., 133, 283, 2005. 101. Abe, N. et al. Zoonotic genotype of Giardia intestinalis detected in a ferret. J. Parasitol., 91, 179, 2005. 102. Homan, W.L. et al. Characterization of Giardia duodenalis by polymerase-chain-reaction fingerprinting. Parasitol. Res., 84, 707, 1998. 103. Matsubayashi, M., Kimata, I. and Abe, N. Identification of genotypes of Giardia intestinalis isolates from a human and calf in Japan. J. Vet. Med. Sci., 67, 337, 2005. 104. van der Giessen, J.W. et al. Genotyping of Giardia in Dutch patients and animals: a phylogenetic analysis of human and animal isolates. Int. J. Parasitol., 36, 849, 2006. 105. Souza, S.L. et al. Molecular identification of Giardia duodenalis isolates from humans, dogs, cats and cattle from the state of Sao Paulo, Brazil, by sequence analysis of fragments of glutamate dehydrogenase (gdh) coding gene. Vet. Parasitol., 149, 258, 2007. 106. Santin, M. et al. Cryptosporidium, Giardia and Enterocytozoon bieneusi in cats from Bogota (Colombia) and genotyping of isolates. Vet. Parasitol., 141, 334, 2006.

Molecular Detection of Foodborne Pathogens 107. O'Handley, R.M. et al. Prevalence and genotypic characterisation of Giardia in dairy calves from Western Australia and Western Canada. Vet. Parasitol., 90, 193, 2000. 108. Trout, J.M., Santin, M. and Fayer, R. Prevalence of Giardia duodenalis genotypes in adult dairy cows. Vet. Parasitol., 147, 205, 2007. 109. Appelbee, A.J. et al. Prevalence and genotyping of Giardia duodenalis from beef calves in Alberta, Canada. Vet. Parasitol., 112, 289, 2003. 110. Mahbubani, M.H. et al. Differentiation of Giardia duodenalis from other Giardia spp. by using polymerase chain reaction and gene probes. J. Clin. Microbiol., 30, 74, 1992. 111. Volotao, A.C. et al. Genotyping of Giardia duodenalis from human and animal samples from Brazil using betagiardin gene: a phylogenetic analysis. Acta Trop., 102, 10, 2007. 112. Lalle, M. et al. Genetic heterogeneity at the beta-giardin locus among human and animal isolates of Giardia duodenalis and identification of potentially zoonotic subgenotypes. Int. J. Parasitol., 35, 207, 2005. 113. Ey, P.L. et al. Comparison of genetic groups determined by molecular and immunological analyses of Giardia isolated from animals and humans in Switzerland and Australia. Parasitol. Res., 82, 52, 1996. 114. Hunt, C.L., Ionas, G. and Brown, T.J. Prevalence and strain differentiation of Giardia intestinalis in calves in the Manawatu and Waikato regions of North Island, New Zealand. Vet. Parasitol., 91, 7, 2000.

51 Isospora

Somchai Jongwutiwes and Chaturong Putaporntip Chulalongkorn University

Contents 51.1. Introduction......................................................................................................................................................................717 51.1.1. Classification of Isospora...................................................................................................................................717 51.1.2. Morphology, Biology, and Life Cycle................................................................................................................718 51.1.3. Epidemiology..................................................................................................................................................... 719 51.1.4. Pathology and Clinical Features........................................................................................................................ 719 51.1.5. Conventional Diagnosis of I. belli..................................................................................................................... 720 51.1.5.1. Sample Collection and Preparation.................................................................................................. 720 51.1.5.2. Conventional Diagnostic Techniques................................................................................................ 721 51.1.6. Molecular Diagnosis of I. belli.......................................................................................................................... 724 51.2. Methods........................................................................................................................................................................... 724 51.2.1. Reagents and Equipment................................................................................................................................... 724 51.2.2. Sample Preparation............................................................................................................................................ 725 51.2.2.1. Disruption of I. belli Oocysts from Stool Samples........................................................................... 725 51.2.2.2. DNA Extraction................................................................................................................................ 726 51.2.3. Detection Procedure.......................................................................................................................................... 727 51.3. Conclusion and Future Perspectives............................................................................................................................... 728 References.................................................................................................................................................................................. 728

51.1  Introduction

51.1.1 Classification of Isospora

The genus Isospora contains at least 14 coccidian species in the phylum Apicomplexa, class Conoidasida, family Eimeriidae. Being one of the most important Isospora species, Isospora belli parasitizes epithelium cells of human small intestines, causing isosporiasis. Historically, description of characteristic oocysts of I. belli in stools was initially observed among military personnel stationed in Egypt, the Middle East and eastern Mediterranean countries during 1914 and 1921.1 Nowadays, it is well recognized that isosporiasis has a cosmopolitan distribution. Although oocysts of I. natalensis were reportedly found in human fecal samples, several studies have failed to reaffirm the presence of these oocysts as a causative agent of diarrheal illness in humans since its first report in1953, raising the possibility that it may not be a valid human pathogenic species.2 Most of the enteric coccidian parasites of man are considered to be opportunistic pathogens except the genus Sarcocystis. Thus, the prevalence of these coccidian infections seems to increase as the number of immunocompromised patients increase, especially after the acquired immunodeficiency syndrome (AIDS) becomes pandemic. In general, severity of illness caused by I. belli seems to be more aggressive in patients with compromised immunity than those with normal immune status. However, chronic emaciating infections have been reported in immunocompetent patients, suggesting that host immunity per se may not determine the severity of isosporiasis.3,4 Meanwhile, differences in the intrinsic virulence of this organism has not yet been explored.

Members of protozoa in phylum Apicomplexa are intracellular organisms that share common structural features by possessing apical complex at certain stages of their life cycles. Apical complex are subcellular organelles consisting of polar ring, rhoptries, micronemes, conoid, and subpellicular microtubules that can be identified under transmission electron microscope. Several lines of evidences indicate that these organelles play a crucial role in host cell invasion and thus exist only at the invasive stages of these protozoa. Parasitic protozoa in the phylum Apicomplexa inhabit tissues or circulation of the host and most of which, including genus Isospora, belong to class Sporozoea. Taxonomic position based on biology and characters of protozoa in subclass Coccidia has placed the genus Isospora in suborder Eimeriina and family Eimeriidae whose members include genus Eimeria and Cyclospora. However, phylogenetic analysis of the small subunit ribosomal RNA sequences as well as internal transcribed spacer 2 region has clustered genus Isospora with tissue-cystforming coccidian in the genus Toxoplasma, Sarcocystis, Hammondia, and Neospora in family Sarcocystidae while non-tissue-cyst forming ones, e.g., Eimeria, Cyclospora, and Cryptosporidium, are in distinct clades.5,6 The presence of tissue cyst in the genus Isospora favors phylogenetic rather than phenetic classification.7–9 Misconception on identification and classification of I. belli over seven decades ago led to a description of I. hominis which de facto belongs to the genus Sarcocystis. Other species of Isospora are parasitic 717

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in wild and domestic animals, e.g., I. arctopitheci in nonhuman primates, I. canis and I. ohioensis in dogs, I. rivolta and I. felis in cats, and I. suis in pigs.10 Over 300 species of Isospora have been proposed and found to infect a wide range of animals, i.e., amphibians, reptiles, birds, and mammals. However, some of these species may require further validation.

51.1.2 Morphology, Biology, and Life Cycle Isospora is an intracellular parasite comprising asexual multiplication and sexual reproduction followed by sporogonic development. The life cycle of I. belli completes within human host without known animal reservoirs. Humans get infected by ingestion of food or water contaminated with oocysts (Figure 51.1). The infective stage of I. belli is characterized by an oval or rugby-shaped oocyst with a wide range of size variation in terms of the maximum length and width, both within and between isolates. The average length measured from more than 700 oocysts is 28.3 μm (range = 17–37). A remarkable narrowing with a neck-like appearance may be observed at one end of some oocysts while most oocysts are nearly symmetry at both ends. The maximum width of oocysts averages 13.5 μm (range = 8–21). The mean shape index or the ratio of length to width of oocyst is 2.1 (range = 1.3–3.3).6 The oocyst wall is bilayer, thin, smooth, and transparent for all stages of development. The inner wall is membranous and the outer wall rigid and relatively impermeable to external fluids.11 When freshly passed in feces, oocyst remains immature, containing a spherical or slightly elongated oval-shaped sporoblast that develops within another rigid cyst wall called sporocyst. Occasionally, some freshly passed oocysts in stool may contain two sporocysts, each of which possesses one sporoblast. The sporocyst of I. belli lacks a proteinaceous

plug, known as a Stieda body. The dimension of sporocyst is 12–14 by 7–9 μm. Sporogony, the process of producing infective sporozoites, of Isospora usually occurs outside the host. The maturation period of oocysts usually completes within 24 h to 10 days.6 The mature oocyst possesses two sporocysts, each with four crescentic or banana-shaped sporozoites clustering together with a clump of granular mass or residual bodies surrounded by sporocyst wall. Detail investigation of ex vivo development has shown that the percentage of oocyst maturation varies between isolates. In vitro studies reveal that only 27–30% of oocysts excreted from stools of patients become fully sporulated at ambient temperature. Importantly, despite the frequently identified characteristic mature oocyst that contains two sporocysts, about 5% of the oocysts that undergo the maturation process are Caryospora-like, characterized by eight sporozoites in a single sporocyst.6,12 Formation of Caryospora-like oocyst requires 5–14 days after excretion in feces.6 Both types of mature oocysts contain a number of granular residual bodies clumping together (Figure 51.2). Environmental factors such as moisture, temperature, and oxygen composition in the atmosphere have been suggested to influence the maturation of Isospora oocysts. It is noteworthy that incubation of I. rivolta oocysts at 50°C for 5 min enhances development of Caryospora-like oocysts and they are infective to mice and cats.13 Sporozoites are released from the oocyst and become free in the lumen of small intestine, especially around distal duodenum and jejunum. Thereafter, sporozoites invade small intestinal epithelial cells where asexual multiplication ensues. All endogenous stages of Isospora reside in parasitophorous vacuoles in the cytoplasm of host cells. Sporozoites of some mammalian Isospora species, e.g., I. suis and I. felis, initially undergo repeated endodyogeny generating type I

Schizogony Merozoites Caryospora-like oocyst with sporozoites

Mature oocyst with sporozoites Developing oocyst

Mature oocyst

Environment Sporogony

Gametogony

Microgametes

Immature oocyst with sporoblasts

Caryospora-like oocyst with sporoblast

Oocyst in stool

Oocyst in lumen

Fertilization

Macrogamete

Figure 51.1  Life cycle of Isospora belli depicting schizogony, gametogony and sporogony. (Illustration by Urassaya Pattanawong.)

719

Isospora

(a)

(b)

(c)

(d)

(e)

Figure 51.2  Oocysts of Isospora belli (a) Oocyst from freshly passed stool containing one sporocyst. (b) and (c) Developing oocysts with two sporoblasts. (d) Mature oocyst with two sporocysts, each with four sporozoites. (e) Caryospora-like oocyst with eight sporozoites in one sporocyst. Scale = 10 μm.

merozoites, in which two daughter cells are generated from a mother cell so that these daughter cells are surrounded by the mother cell membrane.14,15 Repeated endodyogeny probably occurs and generates a number of merozoites. Subsequently, type II merozoites are generated from type I merozoites by schizogonic development in which repeated nuclear division within a single cytoplasmic mass occurs. Cytoplasmic division of multinucleate schizont produces a number of merozoites. Meanwhile, schizogonic development occurs in I. belli but asexual multiplication by endodyogeny has not been identified. Fully developed merozoites disintegrate host cells, invade other epithelial cells and undergo further asexual multiplication cycles. Extraintestinal tissue cysts of I. belli have been found in sections of lymph nodes, spleen, and liver at autopsy of AIDS patients, suggesting that these stages may represent hypnozoites or dormant forms that could be responsible for relapse of symptoms.8,9,16,17 Ultrastructural study has revealed that the extraintestinal tissue cyst is surrounded by a parasitophorous vacuole membrane and characterized by oval or ellipsoidal body measuring about 11 × 8 μm. Conspicuous cyst wall is observed with a thickness varying from 0.7 to 4 μm, enclosing a crescent-shaped zoite. Occasionally, the surface of zoites possesses grooves or small projections. Each zoite contains a single rather than multiple nuclei. The anterior part of zoites possesses polysaccharide-like and lipid-like granules, and apical organelles consisting of rhoptry, microneme, and dense bodies. Although the origin of this developmental process remains elusive, abundant numbers of tissue cysts found at autopsy imply that they probably originated from merozoites, rather than sporozoites from oocyst stage.18 It is noteworthy that extraintestinal stages have been found in feline and canine Isospora species.7 Sexual reproduction or gametogony usually occurs after initiation of asexual cycle for about 1 week. Female gamont develops intracellularly without nuclear division and becomes macrogamete while male gamont undergoes multiple nuclear and cytoplasmic divisions, resulting in production of a number of microgametes. Microgametes disrupt host cells and enter other epithelial cells where macrogametes are located. After fertilization, zygote is formed within the epithelial

cell and sequentially develops into an oocyst. Intraepithelial oocysts containing granular bodies are excreted in feces upon rupture of host cell. The oocysts are resistant to environment including common disinfectants and remain viable in cool and moist condition for months.10

51.1.3 Epidemiology Transmission of I. belli is solely anthroponotic because humans are the only known natural hosts. Before 1935, the prevalence of human isosporiasis seems to be low as only 200 cases were reported.19 However, in certain circumstances when living conditions are overcrowded or sanitation is below the standard level, cases of I. belli infections could be encountered. The majority of infected cases were identified mainly among military or related personnel during 1914–1921.1 Until 1961, when awareness of the infection along with the competency of microscopic detection improved, more than 800 cases were detected in various localities in South America.19 Before the pandemic of HIV infections sporadic or endemic isosporiasis has been reported from several countries both in tropical and temperate zones. The infection rates of I. belli among immunocompetent individuals are usually less than 2% while a remarkable higher rate was observed among those with underlying immunodeficiency. The prevalence of isosporiasis in AIDS patients in Zaire, Haiti, Venezuela, Brazil, and Thailand is 19, 15, 14, 9.9, and 7.6%, respectively.6,20–23 Nevertheless, endemicity, public sanitation, pre-existing anticoccidial drug treatment, and other factors may also affect such prevalence. An 8-year surveillance of intestinal infections among AIDS patients in Los Angeles County revealed the presence of I. belli in 1% of pathogen-positive stool samples.24 Interestingly, most of the I. belli positive samples were from foreign-born patients, especially those from Central America. Travel-related infections may be a contributing factor.

51.1.4  Pathology and Clinical Features Infection with I. belli is usually confined to mucosal epithelial cells of upper small intestine. Endogenous stages are variable in size and structure depending on their growth and development.

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Both schizogonic and sporogonic developmental cycles may persist continuously for an indefinite period if untreated or probably the failure of gut immune response to control the progression of infection. No obvious pathological change is observed in early phase of infection except for some leucocyte infiltration in mucosa and lamina propria, most of which are plasma cells, lymphocytes, and eosinophils.8,25 In chronic isosporiasis, pathological features include shortening or blunting of villi or even atrophy, hypertrophied crypts and lamina propria infiltrated with eosinophils, lymphocytes, and neutrophils.4,26 In some cases, histopathology of isosporiasis may masquerade eosinophilic enteritis.4 However, administration of corticosteroid cannot alleviate the symptoms or even worsen its clinical course. Postmortem examinations reveal that some isosporiasis cases who had underlying AIDS exhibit extraintestinal involvement of lymphoid tissues, causing enlargement of mesenteric lymph nodes, liver, and spleen.8,9,16,17 A spectrum of clinical manifestations of isosporiasis has been noted. Although both immunocompetent and immunocompromised patients are susceptible to I. belli infections, higher prevalence of isosporiasis afflicts the latter more frequently than the former. Incubation period takes about 1 week after ingestion of infective oocysts.27,28 Asymptomatic infection with spontaneous clearance of oocysts is rarely observed. Symptomatic infections with I. belli in immunocompetent hosts present with watery diarrhea, abdominal discomfort, colicky abdominal pain, low grade fever, malaise, anorexia, nausea, vomiting, headache, and dehydration. Experimental infections in human volunteers show that symptoms precede patency of oocysts in stool for a few days. The duration of symptoms continues for about 1 day while oocysts are discharged in stool for more than 1 month.27 Diarrheal episode in some patients can be severe with up to 20 stools per day.29 Diarrheal symptoms usually become chronic if undiagnosed. Chronic infection may turn to malnutrition, steatorrhea, and cachexia. The excretion of oocysts may last for months or years in chronic isosporiasis. A few reports have shown chronic diarrheal symptoms lasting for over a decade in immunocompetent patients.3,4,26 Isosporiasis in immunosuppressed patients, such as those who receive immunosuppressive drugs, prolonged corticosteroid treatment and more commonly in AIDS patients, is generally more severe than those with normal immune status in terms of duration of symptoms and volume of stools. The diarrheal symptoms caused by I. belli seem to be a secretory process that may lead to massive fluid loss, resulting in dehydration and electrolyte imbalance. Fever is frequently observed in dehydrated cases. Co-infections with other enteric pathogens can be detected, especially in AIDS patients.6,21,22,30 Recurrent symptomatic isosporiasis occurs in both immunocompetent and immunodeficient individuals. In a study of isosporiasis in Haitian AIDS patients, relapse occurs in almost half of the cases.21,30 Extraintestinal isosporiasis identified in lymphoid tissues at postmortem in AIDS patients has suggested that these stages could re-invade mucosal epithelial cells and are responsible for clinical relapse.8,9,16,17 Although relapse is believed to be caused by reactivation of these dormant forms, its pathogenesis remains to be elucidated. An

Molecular Detection of Foodborne Pathogens

unusual case of endometrial isosporiasis has been reported in an otherwise immunologically uncompromised host presenting with granulomatous endometritis. Numerous intracytoplasmic cysts resembling I. belli oocysts, but not unicellular zoite, were found in endometrial layer of the uterus.31 Laboratory tests frequently show eosinophilia in a majority of infected cases but remarkable alteration of other hematological profile is not observed. Patients with normal immune status tend to have higher levels of eosinophils than those with suppressed immune system.6 Charcot–Leyden crystals in stools of isosporiasis patients are frequently observed, indicating active proliferation and degradation of eosinophils in gut tissues. Because endogenous stages of I. belli neither cause apparent ulceration of intestinal epithelial cells nor induce intense inflammatory response, no remarkable number of erythrocyte or leucocyte is found in stool. Identification of characteristic oocysts in stool is the definite diagnosis in routine laboratory practice. The efficiency of detection depends on various factors such as microscopist’s experience, shedding pattern of oocysts that may be fluctuating during the course of infection as well as choice of diagnostic techniques. In general, diarrheal symptoms of isosporiasis respond well to treatment with trimethoprim (160 mg) and sulfamethoxazole (800 mg) given orally four times a day for 10 days.21,30 However, almost half of the patients may require long-term prophylactic treatment to prevent relapse. Several anti-relapse regimens have been reported which include trimethoprim plus sulfamethoxazole, pyrimethamine alone or in combination with sulfadoxine.

51.1.5 Conventional Diagnosis of I. belli Conventional laboratory diagnosis of isosporiasis relies on identification of characteristic oocysts in stool or intestinal content. Occasionally, oocysts of I. belli can be overlooked because of its transparency when adjustment of microscope is not appropriate. Although endogenous stages can be observed from tissue biopsy samples, they need to be differentiated from other coccidian, especially those of Cyclospora cayetanensis. Alternatively, molecular techniques can be applied to the detection of I. belli DNA from oocysts in stool or intestinal fluid as well as other endogenous stages in tissue samples. 51.1.5.1 Sample Collection and Preparation Stool sample. Collection of stool samples containing or suspicious of having oocysts of I. belli essentially follows standard guidelines for parasitological diagnosis. Fresh stool samples should be collected without contamination of urine or other materials and kept in a clean container with lid cover to prevent flies or insects that may lay eggs on them. Stool samples should be carefully handled because they may contain various kinds of infectious pathogens, ranging from viruses to parasites. Use universal precautions and wear gloves when handling stool specimens. Although immediate examination of stool specimen is not a crucial issue as those for amoebic trophozoites, it is recommended to examine within 24 h after passage. If examination of stool sample is not possible within

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Isospora

1 day, stool should be kept at less than 10°C, but not frozen, in order to minimize bacterial overgrowth and decay of fecal materials. In case of handling a large number of samples that cannot be processed within a few days, especially during field survey or excessive workload in the laboratory, adding appropriate preservation is mandatory. Preservation of stool samples not only maintains structure of adult worms, ova, trophozoites, and cysts/oocysts, but also halts development of almost all parasite stages. A number of stool fixatives are available, some of which are suitable for specific purposes. Of these, formalin solution in the concentration of 5 or 10% is the most widely used and meets several objectives for subsequent parasitologic examination. It has been recommended that 5% formalin solution is appropriate for preserving protozoan cysts, coccidian oocysts and microsporidian spores while 10% solution is for helminthes, larvae, and eggs. Formalin solution can be diluted with water, phosphate buffer saline or normal saline solution. Preparation of 10% formalin solution is by diluting one volume of the commercially available formalin with nine volumes of appropriate diluents. In fact, the actual concentration of the solution is only 3.7–4% because the commercial stock of formalin is supplied as 37–40% solution. Alternatively, other preservatives that are used infrequently can also be used for fixing stool containing I.  belli oocysts, such as merthiolate–iodine–formalin (MIF) and polyvinyl alcohol (PVA). Usually equal amount of preservative and stool sample are sufficient to retain structure of cysts and ova. However, it is important to mix stool and preservative suspension thoroughly by applicator stick before storage.32 Although coccidian oocysts can be preserved in 5–10% formalin solution for microscopy-based diagnostic purpose, it is not suitable for subsequent biological study or molecular analysis of coccidian parasites. For keeping viability of I. belli oocysts, stool sample should be preserved in 2–3% (weight/ volume) potassium dichromate in water as used for preservation of viable oocysts of Cryptosporidium spp. and Cyclospora cayetanensis.12 When molecular analysis of I. belli oocysts is required, it is recommended to preserve the stool sample in ethanol.6 This is by adding three to four volumes of absolute ethanol to one volume of stool sample and mixing thoroughly to ensure that fecal materials are evenly distributed. Ethanol preserved stool samples can be stored at ambient temperature for subsequent DNA extraction despite the rise of temperature to 40°C in tropical countries.33 Although ethanol is not classified as a parasitologic fixative, preservation of coccidian oocysts in ethanol also gives good results for subsequent acid fast stain. Structures of I. belli and Cryptosporidium oocysts kept in 80% ethanol at ambient temperature for more than 2 years are also well-preserved for acid fast stains.6,33 Duodenal content: Like Giardia lamblia and Strongy­ loides stercoralis, aspiration of duodenal content for direct examination of I. belli can be an adjunctive source other than stool samples. Duodenal content should be submitted to a parasitology laboratory without preservative and examination using wet smear method should be done soon to visualize motility of parasites such as falling leaf pattern of Giardia lamblia trophozoites and rapid sine-curve movement of

Strongyloides stercoralis larvae. Usually nonmotile parasite stages are not vulnerable to time interval between sample collection and microscopic examination. If the sample cannot be examined within a few hours after collection, keeping at less than 10°C is recommended. Preservation of the content with 5–10% formalin solution is adequate for parasitological detection but not for molecular analysis. For molecular diagnosis, centrifugation of the sample prior to preservation in ethanol can reduce the amount of ethanol required. Oocysts of I. belli can be seen by direct smear of duodenal content or acid fast staining procedure. Mucus in the intestinal tract attracts a number of oocysts so that sampling of content containing mucus material may help increasing recovery rate. In case of voluminous content, examination of sediment after centrifugation at 500 × g for 10 min should be done. If the volume of content is small and precludes several preparative diagnostic methods, some authors suggest processing the sample for staining procedure or molecular diagnosis, in stead of preparing for direct wet smear examination. Duodenal content can be obtained from various procedures such as duodenal capsule technique or duodenal aspirate during endoscopic examination of upper gastrointestinal mucosa.34 Histological sample: Biopsy material from upper small intestine during endoscopic examination provides a useful diagnostic sample for I. belli as well as other pathogens that parasitize these regions of the gastrointestinal tract such as Giardia lamblia, Cryptosporidium spp, and Cyclospora cayetanensis. Biopsy material should never be allowed to dry but should be fixed with 10% formalin solution for histopathological study. It is possible to diagnose these parasites in hematoxylin-eosin stained sections under light microscopy. However, sensitivity of the test may be variable because sampling of the region used for biopsy may or may not involve the affected areas. The procedure for hematoxylin-eosin stain as well as preparation of biopsy section is essentially the same as standard pathological method. Various stages of I. belli can be identified in cytoplasm of enterocytes lining upper small intestine while Cryptosporidium spp. are seen as tiny deeply stained bodies at the tips of the mucosal epithelial cells.25,35,36 Biopsy materials are also good sample sources for molecular detection provided that they should kept frozen at less than -20°C without preservatives. 51.1.5.2  Conventional Diagnostic Techniques (i) Stool examination: Direct wet smear: The principle is to sample a small amount of stool or other specimens to search for parasites, ova, trophozoites, cysts, and oocysts. Although characteristic movement of amoebic trophozoites can be observed within 30 min after stool passage, such an immediate attention is not imperative for diagnosis of I. belli oocysts. The procedure is as follows. (1) Add one drop of normal saline solution at the center of a clean glass slide (25 × 75 mm). (2) Use an applicator stick to sample a small amount of formed or soft stool, about 2 mg, and apply to the center of slide.

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(3) Mix and smear thoroughly within a circle of diameter less than 1 cm. (4) Cover with a clean cover-slip (22 × 22 mm). (5) Search for I. belli oocysts under 10 × objective lens by scanning systematically so that all areas of the slide are examined. Confirmation of parasite objects should be done under 40 × objective lens. In case of liquid stool, a drop of normal saline may or may not be required for diluting the sample, depending on the amount of fecal debris. Usually oocysts of I. belli can be best confirmed under 40 × objective lens and the microscope diaphragm should be reduced so that a sharp contrast occurs. The transparency of oocysts may preclude identification under light microscopy if contrast of the microscope field is not properly set. Some microscopists may prefer adding a weak iodine solution to the edge of the cover-slip. Alternatively, a new slide may be prepared by adding one drop of iodine solution instead of normal saline. Concentration by sedimentation: Like most ­helminth eggs and protozoan cysts, oocysts of I. belli can be concentrated by formalin-ether or formalin-ethylacetate sedimentation.37–39 (1) Use about 2–4 g or half a teaspoon of stool specimen and transfer into 10 ml of 5 or 10% formalin solution in a wide mouth container. Use applicator stick to mix the sample thoroughly. Allow the sample to be fixed in about 10 min or longer. If the sample has already been preserved in formalin solution, proceed to the next step. (2) Strain the suspension through wet gauze into a conical 15-ml centrifuge tube. If the specimen contains a lot of mucus, it is not recommended to strain the suspension through wet gauze because the mucus that usually traps a lot of coccidian oocysts may stick to, but not pass through, the gauze. Therefore, the mucus portion should be taken for examination procedure directly. (3) Add 0.85% sodium chloride solution or 5–10% formalin solution to the suspension to make total volume of fecal suspension from step 2 to be 10–12 ml. Centrifuge at 500 × g for 10 min. After decanting out the supernatant, the remaining volume of sediment should be 0.5 or 1 ml. Optionally, this step can be repeated to ensure elimination of excess fecal debris. (4) Add 6–8 ml of 5–10% formalin solution. Total volume of fecal suspension is now about half volume of the tube. (5) Add 3–5 ml of ethyl acetate or diethyl ether. Use a stopper or rubber plug to close the tube. Shake the tube vigorously for at least 30 sec while pressing the stopper or rubber plug with some fingers to counter the effect of vapor pressure in the tube. Leave at room temperature for about 30 sec. Carefully and slowly remove the stopper or rubber plug. (6) Centrifuge at 500 × g for 10 min.

Molecular Detection of Foodborne Pathogens

(7) Gently taking the tube from the centrifuge without disturbing the four layers that should occur in the suspension, comprising in order from top to bottom of the tube: a layer of ethyl acetate (or diethyl ether), a plug of fecal debris, a layer of formalin and a small amount of sediment at the bottom. (8) Remove the plug of debris by ringing the plug with an applicator stick while decant all the supernatant part. Usually a few drops of fluid remain attached to the inner side of the tube and will make a few drops of volume to the sediment. If no fluid remains or the sediment sticks tightly to the bottom of conical tube, a few drops of saline solution or 5–10% formalin solution can be added. (9) Apply one or a few drops of the sediment fluid to a clean glass slide and cover with 22 × 22 mm coverslip. Optionally, one drop of weak iodine solution may be added prior to covering. (10) Examine systematically for the presence of I. belli oocysts as well as other parasites. Concentration by flotation: Besides sedimentation method that can be used to concentrate oocysts of I. belli (and other parasite ova and protozoan cysts), flotation can be an alternative method. Procedure for flotation concentration is originally described by Faust and colleagues.40 The modified procedure by Garcia is as follows.39 (1) Use about 2–4 g or half a teaspoon of stool specimen and transfer into 10 ml of 5 or 10% formalin solution in a wide mouth container. Use applicator stick to mix the sample thoroughly. Allow the sample to be fixed in about 10 min or longer. If the sample has already been preserved in formalin solution, proceed to the next step. (2) Strain the suspension through wet gauze into a roundbottom 15-ml tube. If the specimen contains a lot of mucus, it is not recommended to strain the suspension through wet gauze because the mucus that usually traps a lot of coccidian oocysts may stick to, but not pass through, the gauze. Therefore, the mucus portion should be taken for examination procedure directly. (3) Add 0.85% sodium chloride solution or 5–10% formalin solution to the suspension to make total volume of fecal suspension from step 2 to be 10–12 ml. Centrifuge at 500 × g for 10 min. After decanting out the supernatant, the remaining volume of sediment should be 0.5 or 1 ml. Optionally, this step can be repeated to ensure elimination of excess fecal debris. (4) Re-suspend the sediment with 1 or 2 ml of zinc sulfate (33% aqueous solution, specific gravity 1.18). Add additional zinc sulfate solution to fill the tube within 2–3 mm of the rim. (5) Centrifuge at 500 × g for 2 min. Allow the centrifuge to stop by its own inertia without deceleration or break.

Isospora

(6) Gently taking the tube from the centrifuge without interfering the solution in the tube. Alternatively, some authors recommend not to take the tube out of the centrifuge, but to proceed to the next step while the tube is still in the centrifuge machine. (7) Because the floated materials are on the surface of the solution, harvesting the concentrated samples should be done with care. A Pasteur pipette should be placed exactly at, not below, the surface. Alternatively, a clean wire loop can be used just to touch the surface without dipping into it. (8) Apply the fluid to a clean glass slide and cover with 22 × 22 mm cover-slip. Optionally, one drop of weak iodine solution may be added prior to covering. (9) Examine systematically for the presence of I. belli oocysts as well as other parasites. Acid-fast stain: Conventional microscopy without any staining procedure can detect oocysts of I. belli and has been used in routine diagnostic laboratory. Acid-fast staining method, intentionally deployed for detection of Cryptosporidium spp. and Cyclospora oocysts, has also been found to be useful for diagnosis of I. belli oocysts.6 A number of acid fast staining methods are available. Specimen can be versatile, ranging from fresh stool, sediment from formalin-ethyl acetate sedimentation, ethanol-preserved stool sample, duodenal fluid, or other body fluid. One of these is a modified Kinyoun acid fast stain.41 (1) Make a smear on the clean slide by applying one or two drops of liquid specimen or 2 mg of formed stool. If stool contains mucus, it is recommended to sample the mucus portion because oocysts tend to concentrate in it. Smear should not be too thick. A proper thickness of smear is the one that should be able to see partly through the background. Allow the smear to dry at ambient temperature. (2) Dip the smear into or flood the smear with absolute methanol for 1 min. (3) Stain by dipping the smear into or flooding the slide with Kinyoun’s carbol fuchsin solution. Allow staining time for 1 min. (4) Rinse the stained slide excessively with tap water. (5) Dip or flood the stained slide with 10% sulfuric acid for one or more min until no more color runs from the slide as a decolorization step. (6) Rinse the stained slide excessively with tap water. (7) Counterstain the slide with light green SF yellowish stain for 1 min. (8) Rinse the stained slide thoroughly with tap water. (9) After the slide is completely dry, examine the stained slide with objectives 10 × or 40 ×. If other coccidian oocysts are to be searched for, use objective 100 ×. Autofluorescence: Oocysts of I. belli, Cy. cayetanensis and Cryptosporidium spp. become autofluorescent bright green under violet excitation (405 nm), bluish violet autofluorescent

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under ultraviolet excitation (365 nm) and green autofluorescent under blue-violet light excitation (436 nm).42 Faint reddish fluorescence is also observed under green light (546 nm). Characteristic size, shape, and structure of I. belli oocysts under autofluorescence microscopy differ from those of Cyclospora cayetanensis and Cryptosporidium spp., and thus the method is useful for diagnosis. Autofluorescence is proven to be more sensitive than bright field microscopy in detecting Isospora oocysts.43,44 (1) Prepare stool sample on a clean glass slide following the procedure as described for direct wet smear method. Alternatively, 50–100 μl of unstained stool sediment from concentration method can be used. (2) Examine systematically using 10 × and 40 × objective lens under fluorescent microscope using ultraviolet excitation at 365 nm and/or violet excitation at 405 nm. (ii) Histopathology: Procedures for identification of I. belli in tissue samples essentially follow the standard histophathological method. Biopsy tissue should be fixed with 10% formalin solution, embedded in paraffin or equivalent materials and stain with hematoxylin-eosin. The endogenous stages can be seen in the cytoplasmic mass of tissues or at the lamina propria. Usually the endogenous stages of I. belli are found in duodenum and upper jejunum although involvement of biliary ducts and large intestine has been reported.8,9,16 Although stool examination often gives more sensitive results than histopathology because the sampling areas of biopsy are limited and may not include the affected regions, a large study involving 118 adult AIDS patients who suffered from chronic diarrhea has identified I. belli from duodenal biopsy samples in two patients who had negative results in stool examinations.16 Duodenal mucosal architecture of patients with chronic isosporiasis usually presents with blunting of villi and hypertrophied crypts. Endogenous stages of I. belli in intestinal epitheliums consist of schizogony, gametogony, and early sporogony.25 Inflammatory responses in upper intestinal tissues of both isosporiasis and cyclosporiasis are similar showing mixed infiltration of polymorphonuclear cells, plasma cells, lymphocytes, and eosinophils although the eosinophilic infiltration seems to be more prominent in isosporiasis.25,45 Meanwhile, all endogenous stages of Cryptosporidium spp. are located at intracellular rim at the luminal surface of mucosal epithelium and the diameters of all stages do not exceed 6 μm, making them easily differentiate from I. belli.35,36 Extraintestinal stages of I. belli, existing predominantly in mesenteric lymph nodes, liver, and spleen of some AIDS patients, are mostly crescent-shaped, centrally located and contain only one zoite in tissue cyst. The presence of longitudinal grooves or projections of the zoite surface may cause artefactual findings that two to three zoites are in tissue cysts. Neither multinucleate nor sexual stages exist in tissue cyst of isosporiasis.18

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Molecular Detection of Foodborne Pathogens

51.1.6 Molecular Diagnosis of I. belli Although laboratory diagnosis of I. belli can be performed by searching for characteristic oocysts in stool or other clinical samples under light microscopy, the accuracy of the method largely depends on experienced microscopists. In certain circumstances, endogenous stages of I. belli in tissue samples require differentiation from other intestinal coccidian such as Cy. cayetanensis and Sarcocystis hominis. Because limited genetic data of I. belli is available and microscopy-based method has been used for routine laboratory diagnosis, little progress has been made for the polymerase chain reaction (PCR)-based method to detect I. belli. Amplification of the small subunit ribosomal RNA gene of I. belli by nested PCR has been successfully applied to duodenal biopsy, bile, and stool samples without cross reactivity to other protozoa.6,46 Despite no detail comparative study on

diagnostic value of PCR and microscopy to detect I. belli, nested PCR targeting the small subunit ribosomal RNA gene of I. suis in piglet feces is superior to microscopy.47 Diagnosis of I. belli infection by PCR involves isolation of I. belli DNA from clinical specimens that can be stool, duodenal fluid, tissue sample or other potential sources of infections. Source of I. belli DNA is usually from oocyst stage in stool while tissue sample contains any stages of endogenous development.

51.2  Methods 51.2.1 Reagents and Equipment Reagents and equipment for molecular detection of I. belli are listed in Table 51.1. Note that molecular methodology has been persistently improved and several protocols are

Table 51.1 Reagents and Equipments for Molecular Detection of Isospora Belli Procedure

Reagents

Equipment

Disruption of Oocyst Using glass beads

Glass bead, 425–600 μm (Cat. No. G9268 Sigma, USA) Diethyl ether (optional) TE buffer (10 mM Tris-HCl, pH 7.4 and 1 mM EDTA, pH 8.0) Sterile water (autoclaved)

Adjustable micropipettes 100 μl and 1,000 μl Pipette tips 100 μl and 1,000 μl Microtubes with caps, 1.5 ml Disposable gloves Vortex mixer Microcentrifuge, speed up to 13,000 × g

Freeze thaw

Diethyl ether (optional)

Adjustable micropipettes 100 μl and 1,000 μl

TE buffer (10 mM Tris-HCl, pH 7.4 and 1 mM EDTA, pH 8.0)

Pipette tips 100 μl and 1,000 μl Microtubes with caps, 1.5 ml Cryopreservation tubes, 2 ml Disposable gloves Vortex mixer

Sterile water (autoclaved) Liquid nitrogen

Microcentrifuge, speed up to 13,000 × g Low-temperature resistant container Water bath Metal forceps or metal holder

In-house method

DNA Extraction TE buffer (10 mM Tris-HCl, pH 7.4 and 1 mM EDTA, pH 8.0)

Adjustable micropipettes 100 μl and 1,000 μl

Sterile water (autoclaved)

Pipette tips 100 μl and 1,000 μl

10% Sodium dodecyl sulfate (SDS)

Microtubes with caps, 1.5 ml

Proteinase K (20 mg/ml)

Disposable gloves

Buffer-saturated phenol: chloroform:isoamyl alcohol (25:24:1)

Microcentrifuge, speed up to 13,000 × g

Chloroform

Heat block

Ice-cold absolute ethanol

Refrigerature (–80°C)

Ice-cold 80% ethanol

Dessicator (optional)

3.5 M sodium acetate, pH 5.4 Commercial kit

QIAam DNA Stool Mini kit (Cat. No. 51504) that includes InhibitEX tablet, proteinase K, ASL buffer, AW1 buffer, AW2 buffer, AE buffer, QIAamp spin column and 1.5 ml microtubes

Adjustable micropipettes 10 μl, 100 μl, and 1,000 μl Pipette tips 10 μl, 100 μl, and 1,000 μl Microtubes with caps, 1.5 ml Disposable gloves Microcentrifuge, speed up to 20,000 × g (Continued)

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Isospora

Table 51.1  (Continued) Procedure

Reagents

Equipment Heat block Vortex mixer

DNA amplification

PCR product analysis

Control positive and negative template DNA

Adjustable micropipettes 10 μl and 100 μl

Forward and reverse oligonucleotide primers (0.1 μM each) (see Figure 51.3)

Pipette tips 10 μl and 100 μl

10 × PCR buffer dNTP (2.5 mM each)

Microtubes with caps, 0.2 ml

Taq DNA polymerase or other equivalent enzymes (5 U/μl) Sterile water (Dnase and Rnase free) TBE buffer (0.089 M Tris base; 0.089 M boric acid; 0.002 M EDTA, disodium salt, dihydrate; final pH 8.3) Agarose 5 × gel loading buffer, e.g., 0.25% bromopheol blue; 0.25% xylene cyanol; 15% (w/v) Ficoll type 400 in distilled water

Disposable gloves Microcentrifuge PCR thermal cycler Adjustable micropipettes 10 μl Pipette tips 10 μl Microtubes with caps, 0.2 ml Disposable gloves Horizontal minigel electrophoresis apparatus DC power supply for 50–100 V

Ethidium bromide solution (working concentration is 0.5 μg/ml) DNA marker (100-bp DNA marker ladder)

available. The following procedures can be used as guidelines and definitely can be modified to obtain the best outcomes.

51.2.2 Sample Preparation Freshly collected stool specimen, frozen stool sample or stool preserved in 80% ethanol can be used for extraction of I. belli DNA. Ethanol in preserved stool samples should be removed before disruption of oocysts. Duodenal fluid or tissue samples can be kept at –20°C and ethanol should not be used for preservation of these samples. 51.2.2.1 Disruption of I. belli Oocysts from Stool Samples Because the oocyst wall of coccidian protozoa is solid and resistant to diverse environmental conditions, it is recommended that the oocyst wall should be ruptured so that internal content either sporoblasts or sporozoites expose to the DNA extraction solution. Examples of simple procedures to disrupt oocyst wall of I. belli are mechanical disruption and freeze-thaw method. (i) Mechanical disruption of I. belli oocysts: (1) Transfer approximately 200 mg of formed stool sample or 0.5 ml of liquid stool into 1.5 ml tube with cap. Add 0.5 ml of sterile water. Mix well until stool becomes homogeneously dispersed by vortex. (2) Optionally, add 0.3 ml of diethyl ether. Cap the tube tightly with cap lock and shake vigorously for 3–5 min. (3) Centrifuge at 5,000 × g for 5 min. Discard supernatant.

(4) Add 0.5 ml of TE buffer to the remaining pellet. (5) Mix well using vortex. If the pellet keeps sticking to the bottom of the tube, try mixing vigorously by pipetting. (6) Add 200–300 mg of glass beads, diameter of 425– 600 μm (Cat. No. G9268 Sigma, USA) to the suspension. Shake vigorously for 20 min using vortex mixer. If possible, set at maximum speed. (7) Spin down briefly. (8) Transfer all suspension to a new microcentrifuge tube. Centrifuge at 13,000 × g for 5 min. Discard supernatant and save the pellet for DNA extraction step. (ii) Freeze-thaw method: (1) Transfer approximately 200 mg of formed stool sample or 0.5 ml of liquid stool into 1.5 ml microcentrifuge tube with cap. Add 0.5 ml of sterile water. Mix well until stool becomes homogeneously dispersed by vortex. (2) Optionally, add 0.3 ml of diethyl ether. Cap the tube tightly with cap lock and shake vigorously for 3–5 min. (3) Centrifuge at 5,000 × g for 5 min. Discard supernatant. (4) Add 0.5 ml of TE buffer to the pellet. (If commercial DNA extraction kit will be used in subsequent DNA extraction process, use 0.5 ml of buffer reagent supplied by the kit such as ASL buffer from QIAamp DNA Stool Mini kit). Mix well until the suspension looks homogenous.

726

(5) Transfer the suspension to a 2-ml cryopreservation tube that is resistant to ultralow temperature (such as Nunc CryoTubeTM). (6) Prepare liquid nitrogen in a low-temperature resistant container. Use enough volume to submerge the cryopreservation tube. (7) Set water bath at 95–100°C. (8) Use forceps or metal holder to submerge the cryopreservation tube into liquid nitrogen for 3 min. (9) Immediately heat the cryopreservation tube in boiled water for 3 min. (10) Repeat Steps 7 and 8 for five to seven cycles. (11) Centrifuge at 13,000 × g for 5 min. Discard supernatant and save the pellet for DNA extraction step. (Omit this step when using buffer supplied in commercial kits). 51.2.2.2 DNA Extraction In-house DNA extraction method: This is a standard protocol using proteinase K digestion with phenol-chloroform extraction. (1) Add 440 µl of TE buffer containing 10 mM Tris-HCl, and 1 mM EDTA, pH.8.0, 50 µl of 10% SDS and 10 µl of proteinase K (20 mg/ml) to the pellet prepared from either mechanical disruption or freeze-thaw method as described. Mix well by inverting the tube repeatedly. (2) Incubate at 50°C overnight or at least 4 h. (3) Add equal volume of buffer-saturated phenol: chloroform:isoamyl alcohol (25:24:1). (4) Close the tube tightly and shake vigorously for 5 to 10 min. (5) Centrifuge at 5,000 × g for 5 min at room temperature. Two layers appear, the upper part contains DNA and the lower is solvent (phenol/chloroform/isoamyl alcohol). (6) Gently pipette the upper part of solution to a new tube without disturbing the lower layer and the interface where the whitish layer may appear. (7) Repeat steps 3 to 6 until no visible whitish layer. (8) Add equal volume of chloroform. (9) Shake vigorously for 5–10 min. (10) Centrifuge at 5,000 × g for 5 min. (11) Save the upper layer of solution to the new tube. (12) Add 2.5 to three volume of ice-cold absolute ethanol and 1/10 volume of 3.5 M sodium acetate, pH 5.4. Mix well and keep at –80°C for 1 h. (13) Centrifuge at 13,000 × g for 15 min at 4°C. Allow to stop without using brake. (14) Remove the solution gently without disturbing the DNA pellet at the bottom of the tube by using pipette. (15) Gently rinse with 0.5 ml of ice-cold 80% ethanol. (16) Centrifuge at 13,000 × g for 3 min at 4°C. Allow to stop without using brake.

Molecular Detection of Foodborne Pathogens

(17) Remove the solution gently without disturbing the DNA pellet at the bottom of the tube by using pipette. (18) Dry the sample in desiccators or at room temperature. (19) Re-suspend in 10–20 µl of TE buffer. A remarkable drawback of the conventional method described above is that significant amount of DNA may be lost, especially when a small amount of sample is used. Remnants of enzymatic inhibitors from stool may significantly hinder PCR amplification. DNA extraction using commercial kit: QIAamp DNA Stool Mini Kit (Catalog number.51504, Qiagen, Hilden, Germany) provides a convenient way to extract DNA from I. belli and other biological organisms.48 (1) Add 800 µl of ASL buffer to the suspension prepared from mechanical disruption or add 300 µl of ASL buffer to the sample prepared from freezethaw method. Total volume is now about 800 µl. (2) Add 1 InhibitEX tablet to each sample and vortex immediately and continuously for 2 min or until the tablet is completely suspended. Incubate suspension for 2 min at room temperature. (3) Centrifuge sample at 20,000 × g for 3 min to pellet inhibitors bound to InhibitEX. (4) Pipette all supernatant into a new 1.5 ml tube and discard the pellet. Centrifuge sample at 20,000 × g for 3 min. (5) Pipette 15 µl of proteinase K into a new 1.5 ml tube. (6) Pipette all supernatant from step 4 into a tube of step 5 (Note: if the volume of supernatant is less than 200 µl, add more ASL buffer to make 200 µl in final volume). (7) Add 200 µl AL buffer and mix by vortex for 20 sec. (8) Incubate at 75°C for 20 min. Spin down the sample before next step. (9) Add 200 µl of ethanol (96–100%) to the sample, and mix by vortexing. Spin down the sample before next step. (10) Label the lid of a new QIAamp spin column placed in a 2 ml collection tube. Carefully apply the complete lysate from step 9 to the QIAamp spin column without moistening the rim. Close the cap and centrifuge at 20,000 × g for 1 min. Place the QIAamp spin column in a new 2 ml collection tube, and discard the tube containing the filtrate. (11) Carefully open the QIAamp spin column and add 500 µl AW1 buffer. Centrifuge at 20,000 × g for 1 min. Place the QIAamp spin column in a new 2 ml collection tube, and discard the collection tube containing the filtrate. (12) Carefully open the QIAamp spin column and add 500 µl AW2 buffer. Centrifuge at 20,000 × g for 3 min. Discard the collection tube containing the filtrate.

727

Isospora

(13) Place the QIAamp spin column in a new 1.5-ml microcentrifuge tube. Centrifuge at 20,000 × g for 1 min. Discard the collection tube containing the filtrate. (14) Transfer the QIAamp spin column into a new labeled 1.5 ml microcentrifuge tube and pipette 30–50 µl of AE buffer directly onto the QIAamp membrane. Incubate at room temperature for 5 min, then centrifuge at 20,000 × g for 1 min to elute DNA.

for 30 sec, and 72°C for 1 min; one cycle of 72°C for 5 min. (3) Use 1 µl of primary PCR sample as template DNA for nested PCR. Composition of PCR mix is as follows: Primary PCR product

1.0 µl

Forward PCR primer 3 (0.1 µM)

0.5 µl

Reverse PCR primer 4 (0.1 µM)

0.5 µl

51.2.3  Detection Procedure

10 × PCR buffer (supplied in a PCR kit) dNTP mixture (2.5 mM each)

5.0 µl

We present below a nested PCR targeting the small subunit ribosomal RNA gene for detection of I. belli.6,46 Because of the extreme sensitivity of the nested PCR, it is important to work under clean conditions and use clean equipments to minimize cross contamination and PCR inhibition. Wear clean power-free gloves to handle all the preparative process.

Taq DNA polymerase (5 U/µl) Sterile water (DNase and RNase free)

(1) In a 0.2 ml tube designed to fit a PCR thermal cycler that is pre-cool on ice, mix together: Template DNA

5.0 µl

Forward PCR primer 1 (0.1 µM)

0.5 µl

Reverse PCR primer 2 (0.1 µM)

0.5 µl

10 × PCR buffer (supplied in a PCR kit) dNTP mixture (2.5 mM each)

5.0 µl 4.0 µl 0.5 µl

Taq DNA polymerase (5 U/µl) Sterile water (DNase and RNase free)

34.5 µl

SSU rRNA

IBITS1F0

IBITS1F1

ITS-1

5.8S rRNA

For diagnostic PCR, use positive and negative control DNA in separate reaction tubes. Positive control can be genomic or recombinant DNA of I. belli. Forward and reverse PCR primers 1 and 2 are IBITS1F0 (5′-ATGGACGGTGAGGAAGCGGT-3′) and IBITS 2R0 (5′-CCACCGTATCTCAAAGATGT-3′) whose sequences are derived from the internal transcribed spacers 1 and 2, respectively, of I. belli (Figure 51.3). The amplified product contains 684 base pairs. (2) Place the tube in the preheated PCR thermal cycler and start thermal cycling profile as follows: one cycle of 94°C for 1 min; 40 cycles of 94°C for 30 sec, 60°C

4.0 µl 0.5 µl 38.5 µl

Forward and reverse PCR primers 3 and 4 are IBITS1F1 (5′-ATTCTCTCCCACTTGGTGGAA-3′) and IBITS2R1 (5′-ACACTGCCACACGCGTATT C-3′) whose sequences are derived from the internal transcribed spacers 1 and 2, respectively, of I. belli (Figure 51.3). (4) Place the tube in the preheated PCR thermal cycler and start thermal cycling profile as follows: one cycle of 94°C for 1 min; 30 cycles of 94°C for 30 sec, 60°C for 30 sec, and 72°C for 40 sec; one cycle of 72°C for 5 min. (5) Use 5 µl of the nested PCR sample to check for the presence of amplified DNA products by agarose minigel electrophoresis. The expected PCR product contains 466 bp. The primers derived from internal transcribed spacers 1 and 2 sequences under the PCR conditions as described above give a sensitive and specific result capable of detecting a single oocyst of I. belli. A minimum of five oocysts can be reproducibly amplified by nested PCR and these primers do not cross hybridize with other enteric protozoan as well as human DNA (Figure 51.4). M

1

2

3

4

5

6

7

8

9

10

11 12 13 14 M

466 bp

ITS-2

Primary PCR

Nested PCR

LSU rRNA

IBITS2R0

IBITS2R1

Figure 51.3  Schematic representation of the RNA cistron of Isospora belli. Locations of forward and reverse PCR primers are indicated as directional arrows.

Figure 51.4  Agarose gel electrophoresis of nested PCR amplification using primers derived from internal transcribed spacer 1 (ITS-1) and ITS-2. DNA was extracted by using commercial purification kit (Qiagen). Lanes M, molecular markers of 100-bp DNA ladder; lane 1, positive control; lanes 2 and 4, negative control; lane 3 sample with one oocyst; lanes 5 and 6, samples with five oocysts; lanes 7 and 8, samples with 20 oocysts; lane 9, stool sample from a patient with Isospora belli oocysts in stool; lanes 10–14, Giardia lamblia, Entamoeba histolytica, Cyclospora cayetanensis, Cryptosporidium hominis and human DNA, respectively.

728

51.3 Conclusion and Future Perspectives It has been almost a century ago that I. belli was first recognized as a human pathogen. Since then the prevalence of human isosporiasis seems to be low in general population when comparing with other human enteric pathogenic protozoa. Not until the pandemic of HIV infections, has I. belli been listed as an important opportunistic parasite causing severe and prolonged debilitating diarrheal illness. However, unawareness of this enteric coccidian as a causative agent of diarrhea in immunocompetent hosts as well as lack of diagnostic concern of this pathogen could account for underestimation of the prevalence. Like most other enteric coccidian, developmental cycle of I. belli consists of both asexual and sexual reproductions in a single host. Freshly excreted oocysts from infected human stool are not readily infective but require some time in the environment to reach maturation, producing eight infective crescentic-shape sporozoites per oocyst. The oocysts are environmentally resistant and can retain viability for months at low temperature. Invasion of upper small intestinal enterocytes by sporozoites initiates asexual developmental cycle where schizogony ensues. To date endodyogenic development preceding schizogony has not been identified in I. belli as seen in some other mammalian Isospora species. However, extraintestinal unicellular zoites, found mostly in lymphoid tissues of some AIDS patients, are suggestive of dormant stage that could be responsible for relapse in almost half of symptomatic isosporiasis cases. The high rate of relapse in I. belli infection suggests the presence of drug resistant cryptic stage that is not responsive to treatment. Whether the cryptic stage is an extraintestinal dormant stage and occurs in all I. belli-infected patients require further investigation. Isosporiasis is more prevalent in immunocompromised patients than in immunocompetent individuals and the severity of illness seems to be more in the former. However, chronic emaciating infections have reportedly occurred in patients without compromised immunity, raising the possibility that other factors, in addition to host immunity, such as parasite strain difference could contribute to disease severity. The lack of appropriate animal models hinders experimental studies to elucidate pathogenesis of I. belli. Meanwhile, genomic analysis as well as whole genome sequence determination of I. belli is imperative to shed light on this neglected coccidian. Although conventional stool examinations by direct smear and concentration methods are routinely used to detect I. belli oocysts, the transparency of oocysts can mask correct diagnosis. Staining by acid-fast techniques aiming at diagnosing Cryptosporidium and Cyclospora are also useful to detect oocysts of Isospora but the sensitivity is almost the same as conventional examination by expert microscopist. Alternative diagnosis by examination of wet smear of stool samples by fluorescence microscopy can be used to detect autofluoresed I. belli oocysts. Recent advance in laboratory diagnosis of isosporiasis includes development of PCR-based

Molecular Detection of Foodborne Pathogens

method targeting the ribosomal RNA cistron. Theoretically and practically, the molecular method is far superior to other means of diagnostic techniques. Nevertheless, the presence of PCR inhibitors in stool can drastically lower its sensitivity during DNA extraction process. Addition of recombinant I. belli DNA as a positive control in tested samples can be useful to rule out the presence of such interference. Recent administration of highly active antiretroviral therapy has remarkably reduced the prevalence of several opportunistic infections including isosporiasis. Nevertheless, I. belli can circulate in both immunocompetent and immunocompromised hosts, and has a wide geographic range. The clinical significance of I. belli infection will certainly remain and be in differential diagnosis of diarrheal illness. In this regard, sensitive, and specific diagnostic tools are essential.

References





1. Wenyon, C.M. Coccidiosis of cats and dogs and the status of the Isospora of man. Ann. Trop. Med. Parasitol., 17, 231, 1923. 2. Elsdon-Dew, R. Isospora natalensis (sp. Nov.) in man. J. Trop. Med. Hyg., 56, 149, 1953. 3. Ravenel, J.M., Suggs, J.L. and Legerton, C.W. Human coccidiosis. Recurrent diarrhea of 26 years duration due to Isospora belli: A case report. J. S .Carolina Med. Assoc., 72, 217, 1976. 4. Jongwutiwes, S., Sampatanukul, P. and Putaporntip, C. Recurrent isosporiasis over a decade in an immunocompetent host successfully treated with pyrimethamine. Scand. J. Infect. Dis., 34, 859, 2002. 5. Franzen, C. et al. Taxonomic position of the intestinal protozoan parasite Isospora belli as based on ribosomal RNA sequences. Parasitol. Res., 86, 669, 2000. 6. Jongwutiwes, S. et al. Morphologic and molecular characterization of Isospora belli oocysts from patients in Thailand. Am. J. Trop. Med. Hyg., 77, 107, 2007. 7. Dubey, J.P., and Frenkel, J.K. Extra-intestinal stages of Isospora felis and I. rivolta (Protozoa: Eimeriidae) in cats. J. Protozool., 19, 89, 1972. 8. Restrepo, C., Macher, A.M. and Radany, E.H. Disseminated extraintestinal isosporiasis in a patient with acquired immune deficiency syndrome. Am. J. Clin. Pathol., 87, 536, 1987. 9. Michiels, J.F. et al. Intestinal and extraintestinal Isospora belli infection in an AIDS patient. A second case report. Pathol. Res. Pract., 190, 1089, 1994. 10. Lindsay, D.S., Dubey, J.P. and Blagburn B.L. Biology of Isospora spp. from humans, nonhuman primates, and domestic animals. Clin. Microbiol. Rev., 10, 19, 1997. 11. Beaver, P.C., Jung, R.C. and Cupp, E.W. Coccidia, Microsporidia and Pneumocystis, p. 149–173. In Clinical Parasitology, 9th ed. Beaver, P.C., Jung, R.C. and Cupp, E.W. (eds.). Lea and Febiger, Philadelphia, PA, 1984. 12. Zaman, V. Observations on human Isospora. Trans. R. Soc. Trop. Med. Hyg., 62, 556, 1968. 13. Matsui, T. et al. Infectivity and sporogony of Caryosporatype oocyst of Isospora rivolta obtained by heating. Parasitol. Res., 79, 599, 1993. 14. Ferguson, D.J.P. et al. Ultrastructural observations on multiplication of Cystoisospora (Isospora) felis by endodyogeny. Z. Parasitenkd., 63, 289, 1980. 15. Matuschka, F.R. Ultrastructural evidence of endodyogeny in Isospora suis from pigs. Z. Parasitenkd., 67, 27, 1982.

Isospora 16. Velásquez, J.N. et al. Isosporosis and unizoite tissue cysts in patients with acquired immunodeficiency syndrome. Human Pathol., 32, 500, 2001. 17. Frenkel, J.K. et al. Isospora belli infection: observation of unicellular cysts in mesenteric lymphoid tissues of a Brazilian patient with AIDS and animal inoculation. J. Eukaryot. Microbiol., 50 (Suppl), 682, 2003. 18. Lindsay, D.S. et al. Examination of extraintestinal tissue cysts of Isospora belli. J. Parasitol., 83, 620, 1997. 19. Faust, E.C. et al. Human isosporosis in the Western Hemisphere. Am. J. Trop. Med. Hyg., 10, 343, 1961. 20. Henry, M.C. et al. Parasitological observations of chronic diarrhoea in suspected AIDS adult patients in Kinshasa (Zaire). Trans. R. Soc. Trop. Med. Hyg., 80, 309, 1986. 21. DeHovitz, J.A. et al. Clinical manifestations and therapy of Isospora belli infection in patients with the acquired immunodeficiency syndrome. N. Engl. J. Med., 315, 87, 1986. 22. Certad, G. et al. Isosporiasis in Venezuelan adults infected with human immunodeficiency virus: Clinical characterization. Am. J. Trop. Med. Hyg., 69, 217, 2003. 23. Lainson, R. and da Silva, B.A. Intestinal parasites of some diarrhoeic HIV-seropositive individuals in North Brazil, with particular reference to Isospora belli Wenyon, 1923 and Dientamoeba fragilis Jepps & Dobell, 1918. Mem. Inst. Oswaldo Cruz, 94, 611, 1999. 24. Sorvillo, F.J. et al. Epidemiology of isosporiasis among persons with acquired immunodeficiency syndrome in Los Angeles County. Am. J. Trop. Med. Hyg., 53, 656, 1995. 25. Brandborg, L.L., Goldberg, S.B. and Breidenbach, W.C. Human coccidiosis—a possible cause of malabsorption. N. Engl. J. Med., 283, 1306, 1970. 26. Trier, J.S. et al. Chronic intestinal coccidiosis in man: Intestinal morphology and response to treatment. Gastroenterology, 66, 923, 1974. 27. Matsubayashi, H. and Nozawa, T. Experimental infection of Isospora hominis in man. Am. J. Trop. Med., 28, 633, 1948. 28. McCracken, A.W. Natural and laboratory-acquired infection by Isospora belli. South Med. J., 65, 800, 1972. 29. Syrkis, I. et al. A case of severe human coccidiosis in Israel. Israel J. Med. Sci., 1, 373, 1975. 30. Pape, J.W., Verdier, R.I. and Johnson, W.D. Jr. Treatment and prophylaxis of Isospora belli infection in patients with the acquired immunodeficiency syndrome. N. Engl. J. Med., 320, 1044, 1989. 31. de Otazu, R.D. et al. Endometrial coccidiosis. J. Clin. Pathol., 57, 1104, 2004. 32. Garcia, L.S. Collection, preservation, and shipment of fecal specimens, p. 723–740. In Diagnostic Medical Parasitology, 4th ed. Garcia, L.S. (ed.). ASM Press, Washington, DC, 2001.

729 33. Jongwutiwes, S. et al. Simple method for long-term copropreservation of Cryptosporidium oocysts for morphometric and molecular analysis Trop. Med. Int. Health, 7, 257, 2002. 34. Garcia, L.S. Macroscopic and microscopic examination of fecal specimens, p. 741–785. In Diagnostic Medical Parasitology, 4th ed. Garcia, L.S. (ed.). ASM Press, Washington, DC, 2001. 35. Nime, F.A. et al. Acute enterocolitis in a human being infected with the protozoan Cryptosporidium. Gastroenterology, 70, 592, 1976. 36. Meisel, J.L. et al. Overwhelming watery diarrhea associated with a Cryptosporidium in an immunosuppressed patient. Gastroenterology, 70, 1156, 1976. 37. Ritchie, L. An ether sedimentation technique for routine stool examinations. Bull. U.S. Army Med. Dept., 8, 326, 1948. 38. Young, K.H. et al. Ethyl acetate as a substitute for diethyl ether in the formalin-ether sedimentation technique. J. Clin. Microbiol., 10, 852, 1979. 39. Garcia, L.S. Sputum, aspirates, and biopsy material, p. 809–– 828. In Diagnostic Medical Parasitology, 4th ed. Garcia, L.S. (ed.). ASM Press, Washington, DC, 2001. 40. Faust, E.C. et al. A critical study of clinical laboratory technique for the diagnosis of protozoan cysts and helminth eggs in feces. Am. J. Trop. Med., 18, 169, 1978. 41. Ma, P. and Soave, R. Three-step stool examination for cryptosporidiosis in 10 homosexual men with protracted watery diarrhea. J. Infect. Dis., 147, 824, 1983. 42. Varea, M. et al. Fuchsin fluorescence and autofluorescence in Cryptosporidium, Isospora and Cyclospora oocysts. Int. J. Parasitol., 28, 1881, 1998. 43. Daugschies, A. et al. Autofluorescence microscopy for the detection of nematode eggs and protozoa, in particular Isospora suis, in swine feces. Parasitol. Res., 87, 409, 2001. 44. Bialek, R. et al. Comparison of autofluorescence and iodine staining for detection of Isospora belli in feces. Am. J. Trop. Med. Hyg., 67, 304, 2002. 45. Sun, T. et al. Light and electron microscopic identification of Cyclospora species in the small intestine. Evidence of the presence of asexual life cycle in human host. Am. J. Clin. Pathol., 105, 216, 1996. 46. Müller, A. et al. Detection of Isospora belli by polymerase chain reaction using primers based on small-subunit ribosomal RNA sequences. Eur. J. Clin. Microbiol. Infect. Dis., 19, 631, 2000. 47. Joachim, A. et al. Detection of Isospora suis (Biester and Murray 1934) in piglet faeces--comparison of microscopy and PCR. J. Vet. Med. B, 51, 140, 2004. 48. QIAamp DNA Stool Mini Kit Handbook. For DNA purification from stool samples, August 2001.

52 Sarcocystis

Benjamin M. Rosenthal

United States Department of Agriculture

Contents 52.1. Introduction..................................................................................................................................................................... 731 52.2. Methods........................................................................................................................................................................... 732 52.2.1. Sample Collection and Preparation................................................................................................................... 732 52.2.1.1. Preliminary Visual Screening for Sporocysts.................................................................................. 732 52.2.1.2. Sporocyst Isolation............................................................................................................................ 732 52.2.1.3. DNA Extraction................................................................................................................................ 733 52.2.2. Detection Procedures........................................................................................................................................ 734 52.3. Conclusions and Future Perspectives.............................................................................................................................. 735 References.................................................................................................................................................................................. 736

52.1  Introduction When people eat undercooked beef or pork containing viable Sarcocystis hominis or Sarcocystis suihominis, they can contract acute gastro-intestinal infections that culminate, about two weeks later, with the excretion of parasites infectious for cattle or swine, respectively.1–8 Molecular methods can play a valuable diagnostic role, because neither the sarcocysts (in tissues) nor the oocysts (in feces) are easily distinguished from a large number of other, related parasite species. Other parasites which pose no known risk to human health also form sarcocysts in cattle and swine; and human feces may contain the morphologically similar oocysts of other parasite species. Genetic characterization provides a valuable means to diagnose such infections and can help identify a given infection’s source. By feeding contaminated meat to carnivores, and by feeding oocysts to herbivores, a parasite’s transmission route can be established experimentally. Such methods have established the zoonotic transmission of S. hominis and S. suihominis.8–12 Molecular methods may help identify the cysts that form in intermediate host tissues (sarcocysts), including those of S. hominis in beef and S. suihominis in pork,13–15 but such methods have unfortunately not yet been applied to sarcocysts of human skeletal muscle16–19 or the oocysts in human excreta. Mature, excreted oocysts of S. hominis, S. suihominis, and indeed all species of Sarcocystis, Besnoitia, Frenkelia, Hammondia, and Toxoplasma share the same basic physical attributes: each contains two sporocysts which, in turn, envelop four sporozoites.20 Where the oocyst wall has ruptured, individual sporocysts containing their quartet of sporozoites and granular residuum may be evident. In preparations where the oocyst wall is invisible, parasites will appear as paired sporocysts. An overlapping size range precludes definitive diagnosis of such species using oocyst morphology alone.

Light microscopy provides only marginally more identifying information for sarcocysts in histological sections. For example, a diagnosis of S. hominis can be ruled out when the only sarcocysts present in a beef sample are “thin-walled.” These correspond, instead, to S. cruzi, which employ canine rather than human definitive hosts. Finding “thick-walled” sarcocysts of beef does not establish the occurrence of S. hominis, however, because thick-walled sarcocysts could also represent S. hirsuta, a parasite whose development is completed in cats. The only accepted means of visually differentiating among species of Sarcocystis requires examining of the structure of the sarcocyst wall using transmission electron microscopy.21,22 In particular, the pattern of villar invaginations within the sarcocyst wall provide the basis for morphological diagnosis that has, in turn, served to associate particular genetic sequences with their corresponding parasite species. Because transmission emission microscopy cannot be employed as a routine diagnostic methodology, complementary methods (such as the molecular methods emphasized in this chapter) are useful. Although serological reagents have not been developed for the purpose of diagnosing S. hominis or S. suihominis infections, the efficacy of such an approach has been established for related parasites. Indeed, in the course of developing monoclonal antibodies for the detection of Sarcocystis muris, hybridomas that cross react with S. suihominis and other congeners were identified.23 Efforts to develop serological diagnostic test for bovine neosporosis have sought to demonstrate their specificity by confirming their failure to react with Sarcocystis hominis.24,25 Differentiation among various species of Sarcocystis via isoelectric focusing of total protein extracts has not been developed further since its initial description two decades ago.26 Similarly, although the random amplification of DNA fingerprints using low stringency PCR showed early diagnostic potential,27,28 such methods have not been subsequently elaborated, perhaps 731

732

owing to the vulnerability of such methods to artifacts derived from the amplification of exogenous DNA and the difficulties of replicating such results across laboratories.29 In the last decade, nearly all molecular approaches to the detection and diagnosis of species of Sarcocystis have rested on the amplification and characterization of ribosomal DNA.30–36 Such methods have differentiated among parasites dwelling in the tissues of various livestock (and wildlife) hosts, including those of swine and cattle. For S. hominis and S. suihominis, such molecular methods have been applied to the stages encysted in beef and swine muscle. Although molecular methods have not yet been used to assist the routine diagnosis of human sarcocystosis, preliminary work affirms the efficacy of PCR-RFLP approaches to diagnose S. hominis excreted by human volunteers (Zhao-Qing Yang, personal communication). Such methods demonstrably assist the identification of Sarcocystis excreted by nonhuman hosts, as described below. Confirming, or refuting, suspected cases of human sarcocystosis would be greatly assisted, in the future, by judicious application of molecular diagnostic methods. This chapter reviews how molecular methods have been or could be applied to diagnosing these two foodborne agents, providing an overview of methods to collect and prepare parasites (either from the tissues of intermediate hosts, or from the excreta of definitive hosts), and obtain, characterize, and compare diagnostic sequences. Understanding the frequency and consequences of human exposure to these and related species are goals that can be advanced by obtaining additional sequence data and interpreting them within a broad epidemiological and evolutionary context.

52.2  Methods 52.2.1 Sample Collection and Preparation Noninvasive collection of S. hominis and S. suihominis may be accomplished by proper processing of fecal samples, described below. Exposure to human fecal samples demands strict adherence to universal precautions to safeguard the health and safety of those involved in processing such samples.37,38 52.2.1.1 Preliminary Visual Screening for Sporocysts A modified thick smear procedure frequently employed in the diagnosis of helminthiasis also enables sarcocysts to be visualized from human fecal samples,39 but this procedure’s compatibility with downstream molecular diagnostic procedures has not yet been assessed. A small fecal sample is emulsified in saline and, to exclude fibers and seeds, pressed through a mesh sieve. It is then filtered, transferred to slides, and covered by strips of cellophane measuring 22 × 440 mm that have been soaked for at least 24 h in a solution comprised of 100 ml of glycerin, 100 ml of 6% phenol, and 1 ml of 3% malachite green. After standing for 30 min, these are initially screened at 10 × magnification and then at 40 × to further scrutinize suspected parasites. Using this method, and after confirming suspected positive cases by means of a formalin-ether

Molecular Detection of Foodborne Pathogens

concentration technique, sporocysts were identified in 4.6% and 8.0% of rural residents in the Thai Provinces of Ubon Ratchathani and Khon Kaen, respectively. Other flotation methods (described below) could presumably be effectively paired with this screening method in order to identify and genotype oocysts from populations where infection occurs only infrequently. 52.2.1.2 Sporocyst Isolation Various methods may be employed to enrich fecal samples for coccidian oocysts. The isolation of S. hominis or S. suihominis sporocysts from human feces has rarely been described, and no published accounts are yet available describing the subsequent genotyping of the resulting parasites. Fortunately, methods developed for the excreted stages of other coccidians are undoubtedly applicable to these zoonotic species. Fecal examination of suspected human cases be initiated 14–18 days after ingesting raw or undercooked beef, or 11–13 days after ingesting raw or undercooked pork, reflecting the incubation time that typically precedes the onset of oocyst shedding.1 As is generally true when attempting to identify excreted parasite oocysts,40 it may be necessary to take several samples over the course of a few days before oocysts can be identified, given the intermittent nature of oocyst shedding. Daily fecal examination of three human volunteers identified oocysts of S. hominis no sooner than 10 days, and no later than 40 days post infection.4 Peak shedding occurred on day 18 for one individual and day 14 for the other two. A second study of several individuals reported an average prepatent period of 12±1.8 days and a patent period lasting 8.8±1.1days.41 High density solutions such as those made with sodium chloride, zinc sulfate, sucrose, Percoll, Ficoll-Hypaque concentrate such oocysts on the top surface. These are more successful than formalin-ethyl acetate and related sedimentation methods in preparing oocysts for microscopic inspection1 and also are more conducive to PCR amplification by avoiding formalin-induced cross-linking of DNA to proteins. A typical protocol42 (developed initially for the detection of Cryptosporidium parvum oocysts) suspends the fecal sample in water, then filters this through 50 and 100-mesh wire screens to exclude large, insoluble material. The resulting solution is concentrated via centrifugation at 550 × g for 7 min. If prolonged oocyst storage is desired, these can be resuspended in 2% w/v potassium dichromate solution and held at 4°C. Potassium dichromate can later be removed by successive rounds of centrifugation in an excess of distilled water, after which oocysts can be forced to the surface by increasing the specific gravity of the solution as follows: Mix 1 ml of the sporocyst suspension thoroughly with 1 ml of a solution prepared by dissolving 128 g sucrose in 100 ml of water (resulting in a solution with a specific gravity of 1.266). Then, add more sucrose solution until the meniscus is readily accessible. Sporocysts can be obtained by pipetting or touching a cover slip to the surface. A similar approach for obtaining sporocysts of S. neurona from the feces and mucosal scrapings of opossums for the purpose of genetic characterization.43 Here, suspensions are

Sarcocystis

mixed with an equal volume of 5% bleach (NaOCl), incubated on ice for 30 min and stirred every 10 min. Gauze mesh filtration replaces the metal sieves described above, and sporocysts are washed at least three times in fresh saline solution after pelleting the sporocysts using 10 min centrifugation at 2,000 × g. Washing is deemed complete when the odor of bleach is no longer detectable. Sporocysts are then stored at 4°C in Hanks’ buffered salt solution (HBBS) with 100 IU/ml penicillin G and 100 μg/ml streptomycin to discourage the growth of bacterial and fungal contaminants. Further purification can be accomplished by centrifuging such preparations for 10 min at 2,000 × g, resuspending the pellet in 1 ml HBSS, adding this to a 20/30/60% Percoll step gradient, and centrifuging them in a swing rotor at 22,500 × g for 30 min at 25°C. Sporocysts thereby concentrate at the 30/60% Percoll interface and may be subsequently washed by repeated centrifugation at 2,000 × g in HBSS. Most bacterial contaminants are thereby removed. Faster screening methods are appropriate when searching many samples for the rare occurrence of oocysts in ­population-based surveys. For example, an ambitious national survey seeking to estimate the prevalence of viable T. gondii (and other tissue cyst-forming coccidia) in retail meats involved daily fecal assays from thousands of cats that had been fed samples of beef, chicken, or pork44 using modifications of previously described methods45 Fecal samples (30–65 g) were first emulsified in water and a 5–10 g portion was then mixed with a 2 M sucrose solution, filtered through gauze, and centrifuged in 50-ml tubes for 10 min at 1,200 × g. From the surface, a few drops were examined microscopically for oocysts. The top 1 ml was mixed with 10 ml of 2% sulfuric acid in a 50-ml tube in the event that preliminary inspection suggested the need for follow up. Similar methods were used in a successful surveys of coccidian oocysts in thousands of fecal specimens derived from dogs and cats.46,47 Using a floatation solution of 262 mg/ml ZnCl2 and 275 mg/ml NaCl, floated material is transferred to a slide and examined by light microscopy using a magnification of at least 200 × . Positive fecal samples can then be re-examined by a combined sedimentation and flotation procedure. At least 200 ml of tap water is added for each 10 g of sample. The mixture is sieved using a strainer (0.5–0.8 mm mesh aperture) and sedimented for at least 1 h. At most 5 ml of sediment is transferred to a 50-ml tube, mixed with 50 ml concentrated sucrose (550 g sucrose in 450 ml tap water; specific gravity 1.3) and centrifuged at 1,600 × g for 10 min. (A few drops may be examined from the surface via microscopy at this point). For oocyst isolation, 2 ml tap water is gently pipetted on top of the supernatant and the floated material above the sucrose solution is suspended by gently stirring the water and then transferring to a fresh 50 ml tube. Three successive washes in tap water via centrifugation (1,600 × g, 10 min, without brake) and resuspension is followed by final addition of K2Cr2O7 to a final concentration of 1–2% (w/v). To identify and process oocysts of Neospora caninum from dogs feces, which are unlikely to contain either seeds

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or plant fibers, employed no sieving or filtration steps may be necessary.48 A fecal slurry, repeatedly mixed with 2 volumes of water at 37°C, centrifuged at 2,000 × g for 15 min, and resuspended in water, will eventually clear. The resulting pellet is resuspended in one volume of water and mixed with one volume of Sheather’s sugar solution for oocyst flotation. Such oocysts are then washed twice with water, twice with 0.05% aqueous Triton-X 100, twice again with water. This oocyst pellet is resuspended in 2.62% aqueous bleach for five min, centrifuged for 10 min at 2,000 × g, and washed another five to six times prior to DNA extraction (described below). 52.2.1.3 DNA Extraction The oocyst and sporocyst walls represent formidable barriers to accessing the nucleic acids residing within sporozoites. This section begins by reviewing a single tube procedure that has been successfully used to liberate amplifiable DNA from the oocysts of various coccidians. The more traditional approach is then described, employing successive steps to break the oocyst wall, induce the sporozoites to exit their sporocysts, and ultimately extract the DNA via lysis of the sporozoite membrane. (i) Single tube extraction from oocysts: A simple, singletube procedure for extracting PCR-amplifiable DNA from coccidian oocysts has been described that promises good yields using relatively straightforward procedures.49 Cysts of Eimeria nieschulzi were employed in the published account comparing this approach to alternatives, but it has reportedly also been successfully applied to the cysts of Hammondia, Neospora, Isospora, and Caryospora.49,50 Future efforts to extract DNA from the sporocysts of either S. hominis or S. suihominis should consider this approach because it requires relatively little time, yields abundant DNA, and supports the enzymatic amplification of large gene fragments. The initial report of this method succeeded in amplifying nuclear 18S rDNA from as few as ten oocysts, whereas 50 oocysts were required to amplify a 23S rDNA encoded in the plastid genome. In a continuous incubation at 65°C, this approach strips away both the thick outer oocyst layer, comprised of chitin and quinone-tanned protein, and the inner oocyst layer, comprised of lipids and proteins. Briefly, proteinase K digests, detergents dissolve, and bleach strips away the outer layer. High concentrations of EDTA, N-lauroylsarcosine, and proteinase K then dissolve the inner oocyst wall, sporocyst walls and sporozoite membranes. CTAB is included to remove polysaccharides that might bind and inhibit the enzymatic amplification of DNA. To prepare the oocysts for lysis, wash them repeatedly by centrifugation if they have been stored in 2% K 2Cr2O7 solution (i.e., four times at 1.4 × 103 rpm for 5 min) in sterile, high-salinity, phosphate-buffered saline (300 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.7 mM NaH2PO4). Resuspended the oocyst pellet in 200 μl 5.75% sodium hypochlorite and incubate on ice for 30 min. Dilute this oocyst suspension with 1ml sterile ddH2O, pellet by centrifugation, and wash in the saline solution three times to remove the bleach.

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Lysis is accomplished by resuspending these oocysts and incubating them at 65°C for 45 min in 60 μl of a buffer comprised of 660 mM EDTA, 1.3% N-lauroylsarcosine, 2 mg/ ml proteinase K, pH 9.5. Then add 350 μl cetyl–trimethyl ammonium bromide (CTAB) buffer (2% w/v CTAB, 1.4 M NaCl, 0.2% β-mercapto-ethanol, 20 mM EDTA, 100 mM Tris) and incubate for 1 h at 60°C. DNA can then be extracted from the lysate using a traditional phenol/chloroform procedure designed to separate the water-soluble constituents (including nucleic acids) from lipids and most proteins. (Care must be exercised when using and disposing of such organic solvents, and it may be possible to substitute alternative extraction procedures to avoid their use). The published protocol continues by adding an equal volume of phenol/chloroform/isoamyl alcohol (25:24:1) to the lysate. Gently vortex this mixture. Resolve the resulting cloudy suspension into discrete layers by brief centrifugation. Remove the top, aqueous layer being careful not to carry over organic material situated at the tube’s bottom, or at the white interface that may have formed. Any such carry over can be removed by repeating the steps described in this paragraph. To the aqueous solution, add 0.04 volumes of 4 M NaCl and 2–3 volumes of 100% cold ethanol in order to precipitate the DNA. (Glycogen may be added at this step as a co-­precipitant in order to increase the size and visibility of the pellet; it should not interfere with downstream applications). After 20–30 min incubation at −20°C, pellet the DNA by centrifugation. Decant the supernatant and wash with 70% ethanol, taking care not to dislodge the DNA pellet. Air-dry the DNA pellet to remove the ethanol, and resuspend it in sterile water or TE buffer (10 mM Tris, pH 8.0, 1 mM EDTA pH 8.0). More traditionally, a series of steps have been employed to access the DNA residing in the nuclei of sporozoites bounded by sporocyst and oocyst walls. Various means have been employed to breach these barriers, including sonication, agitation in the presence of beads, and freezing in liquid nitrogen and grinding. After liberating sporocysts from their oocyst walls, most procedures then seek to emulate conditions within the gut of the intermediate host that induce sporozoites to exit the oocyst (excyst). As reports describing the application of such in vitro excystation procedures to either S. hominis or S. homini are lacking, this chapter will review how this challenge has been addressed for related parasite species. (ii) In vitro excystation method: Excystation procedures have been described in useful detail for the oocysts of several species of Sarcocystis that establish infections when ingested by caprine and ovine hosts.51 Starting with oocysts stored at 4°C in water containing 20 IU/100 ml of both penicillin and streptomycin, and 100 ug/100 ml amphotericin B, the method calls for 20 min preincubation of sporocysts in a 6–8% bleach solution (NaOCl) at room temperature. (Notably, excystation requires pre-exposure to bleach, but overexposure may harm the resulting sporozoites; thus, 20 min NaOCl pre-incubation are preferable to 10 min pre-incubations, but incubations of over 1 h are inadvisable). Five successive washes in distilled water are then used to remove the bleach. Greatest success can be achieved by resuspending such sporocysts in 1.5 ml

Molecular Detection of Foodborne Pathogens

RPMI 1640 cell culture medium, placing them in an ice water bath, and sonication them for 2 min on 30% duty cycle and output control 3 using a double step micro tip. A final, hour-long incubation (at 39°C in a solution of RPMI 1640 medium, 10% fetal calf serum, and 15% pooled bovine bile) completes the procedure, and largely overcomes the reduction in excystation otherwise observed if the sonication step is skipped. Incubating sporocysts of S. neurona overnight at 37°C and 5% atmospheric carbon dioxide in horse bile containing 2% trypsin induces the excystation of Sarcocystis neurona.43 Considering that raccoons, armadillos, cats, and other mammals may also serve as intermediate hosts of this parasite, the chemical cues for natural excystation are evidently not exclusive to the bile of horses. Physical disruption facilitates the extraction of DNA from intact oocysts that have been concentrated, washed in detergent, and incubated in bleach as described above.48 After removing all residual bleach by pelletting the oocysts in fresh aliquots of water, the oocysts are resuspended in an extraction buffer comprised of 50 mM Tris, pH 8.0, 100 mM NaCl, 50 mM EDTA, 2% SDS and placed in a 50 ml flask containing glass beads 2 mm in diameter. After stirring these for 10 min at room temperature, Proteinase K is added to a final concentration of 1% w/v and incubated at 65°C for 2 h. The procedure is completed with a phenol:chloroform:isoamyl alcohol extraction similar to the one described above. (iii) Extraction from sarcocysts and cell cultures: Genomic DNA is more readily obtained from parasites encysted in the muscle of intermediate hosts because the sarcocyst wall is readily breached by conventional lysis buffers. Hence, commercial kits designed for isolating DNA from animal tissues, for example Qiagen’s DNAeasy columns, routinely suffice for this purpose. Naturally, the resulting extracts will include an abundance of host DNA, requiring that care be taken to selectively amplify only targets of parasite origin. That can be accomplished either by amplifying partial sequences whose termini each differ markedly between coccidia and their vertebrate hosts (as described below) or by obtaining full-length sequences by concatenating overlapping sequences derived from PCR pairing a primer targeting motifs universally conserved among eukaryotes with a primer specifying sequences unique to the coccidian.13 Similar approaches can be successful when parasites have been established in cell cultures.52,53

52.2.2  Detection Procedures Sequence variation in the 18S rDNA (and, to a lesser extent, the adjacent ITS and LSU rDNA subunits) provide the principal molecular basis for discriminating among species of Sarcocystis,15,36,54 for elucidating interrelationships among them,55–63 and for comparing their extent of intraspecific diversity.13 When parasite taxa are closely related, i.e., S. neurona and S. falcatula, the 18S rDNA may not have accumulated an appreciable extent of differentiation, necessitating the search for markers sufficiently variable to provide

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Sarcocystis

resolution about more recent evolutionary events.43,64–68 Furthermore, the possibility exists that very small differences may exist among duplicated (paralagous) copies of the gene within a given genome, a phenomenon which has been more thoroughly established in other Apicomplexan species.69–71 In spite of these caveats, the 18S rDNA remains the molecule of choice for systematics and diagnostics for three important reasons: (i) very highly conserved portions of the molecule flank more variable domains, facilitating PCR amplification of diagnostic regions; (ii) the molecule occurs repeatedly in the parasite genome, increasing the number of PCR targets; and (iii) the molecule has been characterized in many parasite species, establishing a needed frame of reference for identifying unknown specimens. Several methods are available for exploiting such diagnostic variation. The most comprehensive approach is to sequence the complete rDNA from parasites. Such sequences should first be confirmed as having derived from a species of Sarcocystis (as opposed to, say, the host or to other potential contaminants) by querying nucleotide databases for homologous sequences (i.e., a BLAST query of the non-redundant nucleotide database of GenBank). Thereafter, newly acquired sequences can be profitably compared to many related sequences in order to establish their likely identity, using multiple sequence alignment and phylogenetic reconstruction methods. The resulting gene tree establishes whether or not a given sequence corresponds, exclusively, to S. hominis, S. suihominis, or to any other previously characterized taxon. Sequences for which no clear precedent exists enlarge our understanding of parasite biodiversity.72,73 The 18S rDNA need not necessarily be sequenced in its entirety in order to differentiate among known alternatives. Several successful studies have illustrated the diagnostic information in partial rDNA sequences, characterized either by sequencing or by mapping the location of restriction endonuclease polymorphisms15,74–76 Especially where the sampling scope is great or the available resources are limited, such efficiencies offer attractive alternatives to complete sequencing. For the purpose of discriminating among well-defined alternatives, restriction endonuclease digestion of PCR products can be useful. Accordingly, an approximately 900 bp fragment of 18S rDNA has been used in several studies, amplified by primers 2L (5′-GGA TAA CCT GGT AAT TCT ATG-3′) in conjunction with primer 3H (5′-GGC AAA TGC TTT CGC AGT AG-3′). For several species of Sarcocystis, including S. hominis and S. suihominis, exemplary sequences of this gene are available in GenBank, and the digestion patterns of several amplification products, when incubated with any of 12 restriction endonucleases, are known.76 Other available means to characterize single nucleotide polymorphisms77–80 have not yet been applied to problems in Sarcocystis diagnosis or epidemiology. We present below a step-wise protocol for molecular identification of S. hominis and S. suihominis based on PCR amplification of a 915-bp fragment from Sarcocystis 18S rDNA gene using primers 2L and 3H followed by DNA sequencing analysis as reported by Yang et al.76

Procedure: (1) Prepare PCR mixture (50 μl) containing 20 mM Tris–HCl, pH 8.3, 60 mM KCl, 2 mM MgCL2, 10 pM of each primer (2L: 5′-GGA TAA CCT GGT AAT TCT ATG-3′ and 3H: 5′-GGC AAA TGC TTT CGC AGT AG-3′), 200 mM dNTPs and 1.25 U Taq polymerase, 2–4 μl of purified DNA and distilled water (to 50 μl). (2) Carry out PCR amplification on a thermocycler using the following cycling programs: one cycle of 95°C for 2 min; 40 cycles of 94°C for 40 sec, 55°C for 30 sec or 1 min, 72°C for 1 min; one cycle of 72°C for 6 min. (3) Analyze the PCR products by 1% agarose gel electrophoresis containing 0.5 μg/ml ethidium bromide and purify the fragments using a commercial kit (or other technique). (4) Determine the nucleotide sequence of the purified PCR products with the same primers as for PCR using the BigDye™ terminator cycle sequencing kit (PE Biosystems, Foster City, CA) on an ABI 377 DNA automatic sequencer. (5) Align DNA sequences with those available from GenBank with DNASTAR (DNASTAR Inc); undertake homology and divergence analysis with DNASTAR Megalign; perform distance analysis, Neighbor-Joining (N-J) method with PAUP version 4.0. Note: The expected size of PCR product from Sarcocystis 18S rRNA gene using primers 2L and 3H is 915 bp.76 Nucleotide homology in this 18S rRNA gene region is quite high within species (90.0–99.9%), but it is much lower among the different species (71.1–90.7%). Similarly, nucleotide divergence in this 18S rRNA gene region is 0.1–0.6% (mean 0.45) within same species, but it is 1.5–8.6% (mean 3.2) among different species.76

52.3 Conclusions and Future Perspectives As reviewed elsewhere in this book, coccidian parasites exploit two fundamental means of transmission. The first are directly transmitted among the members of a single host species via a fecal-oral route. Fecal contamination of drinking or irrigation water (and therefore, of food) can sustain transmission of such “one-host” coccidia. These parasites are evolutionary related to one another and consequently share many morphological, epidemiological, and physiological attributes. Indeed, these parasites share much in common in spite of their historical segregation into various genera, including Isospora belli and Cyclospora cayetenensis (discussed elsewhere in this book) and the hundreds of species assigned to the genus Eimeria, including several parasites of poultry responsible for significant economic losses. Molecular phylogenetics, based primarily upon ribosomal DNA sequencing, underscore their

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shared common history (and also illustrate their fundamental dissimilarity from species of Cryptosporidium).81 Such methods have enhanced our understanding of the diversity, hostspecificity, and interrelationships among such parasites. These methods have also been successfully applied to diagnosing and studying parasites in the other major group of coccidia: those acquired by carnivores not by imbibing contaminated water, but rather by eating prey animals in whose tissues parasites have encysted. In place of the direct transmission described above, these parasites alternate between herbivores (infected by excreted oocysts) and carnivores (infected by encysted sarcocysts). Although important and interesting distinctions delineate particular members of this group, such “tissue-cyst forming” coccidia share a common evolutionary origin and therefore share many biological attributes. Thus, Sarcocystis hominis and S. suihominis represent but two of hundreds of named species in the Sarcocystidae, which also includes parasites variously assigned to the genera Toxoplasma, Besnoitia, Frenkelia, Hammondia, Neospora, and Sarcocystis. For every study of S. hominis or S. suihominis, more than 100 have been published for Toxoplasma gondii owing to its especially great clinical significance as a major threat to human reproductive health and a serious opportunistic pathogen of AIDS. This justifiably disproportionate research effort has resulted in a suite of experimental methods and assays, including tools of modern molecular genetics, immunological reagents, in vitro culture protocols, in vivo phenotyping, genome sequences, population genetics, expression analysis, and more. Therefore, the broader literature on parasites belonging to the Sarcocystidae, in general, and on T. gondii, in particular, are appropriate resources where the limited literature on S. hominis or S. suihominis is found to be lacking. Having established that eating undercooked beef or pork invites the risk of contracting gastrointestinal parasites whose cysts infectious for livestock, might we wonder whether consuming other livestock or wild game exposes us to as-yet undescribed parasites? Such speculation should be constrained by the understanding that most human pathogens were acquired either from other primates, with whom we share much physiologically, or from our livestock, with whom we have especially close and sustained contact.82 This general insight clearly applies to Sarcocystis spp., where the two known zoonotic species derive from livestock and where non-human primates are competent infection reservoirs.8,83 Accordingly, the zoonotic potential of additional species Sarcocystis should be investigated from other food animals and from any animals also preyed upon by primates that are either strictly carnivorous (i.e., tarsiers) or omnivorous (i.e., baboons and most especially chimpanzees, our closest living relative and our most frequent partner in pathogen exchange). The potential for atypical hosts to sustain infections upon newfound exposure is illustrated by reports that zoo animals (i.e., bison, antelopes) are vulnerable to species of Sarcocystis typical of locally endemic hosts,32,84,85 by serendipitous reports of infections of captive primates with previously unknown species of parasites,3 and by growing recognition that certain

Molecular Detection of Foodborne Pathogens

coccidian parasites occupy decidedly broad host ranges.45,86,87 Serological surveys document further exposure of primates to coccidian parasites, although exposure may not necessarily result in the establishment of infection.88 Similarly, the extent of our vulnerability to cysts excreted by other carnivores remains unclear. Human toxoplasmosis can be contracted by ingesting oocysts excreted by cats. Assorted case reports and surveys of autopsied tissues affirm human exposure to species of Sarcocystis of uncertain provenance.16–18 Subjecting such material to rDNA sequencing would assist in understanding the source of such infections. A previously identified parasite might thereby be recognized as possessing heretofore unknown zoonotic potential if the resulting sequence perfectly matched a previous record. Alternatively, a sequence only imperfectly resembling known homologues might provide valuable clues as to likely candidates nonhuman hosts, as ancestral raptors, felids, canids, vipers, and marsupials each harbored parasites which have subsequently given rise to distinct clades of extant parasite species.57,89–94 In conclusion, molecular methods can and should be employed to aid diagnosis of human infection with S. ­hominis and S. suihominis. Such methods, moreover, offer great potential in exploring the evolutionary origins and exposure routes of other human parasites whose identity and provenance remains enigmatic. Taking advantage of a vast repertoire of methodologies already established in Toxoplasma gondii, and taking into account the growing body of comparative sequence data, should lead to ­important future insights concerning parasite ecology, epidemiology, and evolution.

References



1. Fayer, R. Sarcocystis spp. in human infections. Clin. Microbiol. Rev., 17, 894, 2004. 2. Chen, X.W., Zuo, Y.X. and Hu, J.J. Experimental Sarcocystis hominis infection in a water buffalo (Bubalus bubalis). J. Parasitol., 89, 393, 2003. 3. Gozalo, A.S. et al. Chronic polymyositis associated with disseminated Sarcocystosis in a captive-born rhesus macaque. Vet. Pathol., 44, 695, 2007. 4. Chen, X., Zuo, Y. and Zuo, W. [Observation on the clinical symptoms and sporocyst excretion in human volunteers experimentally infected with Sarcocystis hominis]. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi, 17, 25, 1999. 5. Lian, Z. et al. [Studies on man-cattle-man infection cycle of Sarcocystis hominis in Yunnan]. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi, 8, 50, 1990. 6. Bunyaratvej, S., Bunyawongwiroj, P. and Nitiyanant, P. Human intestinal sarcosporidiosis: report of six cases. Am. J. Trop. Med. Hyg., 31, 36, 1982. 7. Li, J.H. et al. [Sarcocystis suihominis infection in human and pig population in Guangxi]. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi, 22, 82, 2004. 8. Fayer, R. et al. Transmission of Sarcocystis suihominis from humans to swine to nonhuman primates (Pan troglodytes, Macaca mulatta, Macaca irus). Zbl. Parasit., 59, 15, 1979. 9. Dubey, J.P., Fayer, R. and Speer, C.A. Experimental Sarcocystis hominis infection in cattle: lesions and ultrastructure of sarcocysts. J. Parasitol., 74, 875, 1988.

Sarcocystis 10. Li, J.H. et al. [Experimental infection of Sarcocystis suihominis in pig and human volunteer in Guangxi]. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi, 25, 466, 2007. 11. Piekarski, G. et al. [Clinical, parasitological and serological investigations in sarcosporidiosis (Sarcocystis suihominis) of man (author’s transl)]. Immun. Infekt., 6, 153, 1978. 12. Heydorn, A.O. [Life-cycle of Sarcosporidia. IX. Developmental cyclus of Sarcocystis suihominis n. spec.]. Berl. Munch. Tierarztl. Wochenschr., 90, 218, 1977. 13. Rosenthal, B.M., Dunams, D.B. and Pritt, B. Restricted genetic diversity in the ubiquitous cattle parasite, Sarcocystis cruzi. Infect. Genet. Evol., 8, 588, 2008. 14. Vangeel, L. et al. Molecular-based identification of Sarcocystis hominis in Belgian minced beef. J. Food Prot., 70, 1523, 2007. 15. Gonzalez, L.M. et al. Differential molecular identification of Taeniid spp. and Sarcocystis spp. cysts isolated from infected pigs and cattle. Vet. Parasitol., 142, 95, 2006. 16. Wong, K.T. and Pathmanathan, R. Ultrastructure of the human skeletal muscle sarcocyst. J. Parasitol., 80, 327, 1994. 17. Wong, K.T. et al. Light microscopic and three-dimensional morphology of the human muscular sarcocyst. Parasitol. Res., 80, 138, 1994. 18. Pathmanathan, R. and Kan, S.P. Three cases of human Sarcocystis infection with a review of human muscular sarcocystosis in Malaysia. Trop. Geogr. Med., 44, 102, 1992. 19. Mehrotra, R. et al. Diagnosis of human Sarcocystis infection from biopsies of the skeletal muscle. Pathology, 28, 281, 1996. 20. Gardiner, C.H., Fayer, R. and Dubey, J.P. An atlas of rotozoan parasites in animal tissues. Armed Forces Institute of Pathology, American Registry of Pathology, Washington, DC, 1998. 21. Dubey, J.P., Fayer, R. and Speer, C.A. Sarcocystosis of animals and man. CRC, Boca Raton, FL, 1989. 22. Dubey, J.P., Speer, C.A. and Charleston, W.A. Ultrastructural differentiation between sarcocysts of Sarcocystis hirsuta and Sarcocystis hominis. Vet. Parasitol., 34, 153, 1989. 23. Tenter, A.M. et al. Detection of species-specific and crossreactive epitopes in Sarcocystis cystozoites by monoclonal antibodies. Parasitol. Res., 77, 212, 1991. 24. Baszler, T.V. et al. Serological diagnosis of bovine neosporosis by Neospora caninum monoclonal antibody-based competitive inhibition enzyme-linked immunosorbent assay. J. Clin. Microbiol., 34, 1423, 1996. 25. Lally, N.C., Jenkins, M.C. and Dubey, J.P. Evaluation of two Neospora caninum recombinant antigens for use in an enzymelinked immunosorbent assay for the diagnosis of bovine neosporosis. Clin. Diagn. Lab. Immunol., 3, 275, 1996. 26. Tenter, A.M., Johnson, M.R. and Zimmerman, G.L. Differentiation of Sarcocystis species in European sheep by isoelectric focusing. Parasitol. Res., 76, 107, 1989. 27. Joachim, A. et al. A RAPD-PCR derived marker can differentiate between pathogenic and non-pathogenic Sarcocystis species of sheep. Mol. Cell. Probes, 10, 165, 1996. 28. Granstrom, D.E. et al. Differentiation of Sarcocystis neurona from eight related coccidia by random amplified polymorphic DNA assay. Mol. Cell. Probes, 8, 353, 1994. 29. Penner, G.A. et al. Reproducibility of random amplified polymorphic DNA (RAPD) analysis among laboratories. Genome Res., 2, 341, 1993. 30. Heckeroth, A.R. and Tenter, A.M. Development and validation of species-specific nested PCRs for diagnosis of acute sarcocystiosis in sheep. Int. J. Parasitol., 29, 1331, 1999. 31. Tenter, A.M., Luton, K. and Johnson, A.M. Species-specific identification of Sarcocystis and Toxoplasma by PCR amplification of small subunit ribosomal RNA gene fragments. Appl. Parasitol., 35, 173, 1994.

737 32. Fischer, S. and Odening, K. Characterization of bovine Sarcocystis species by analysis of their 18S ribosomal DNA sequences. J. Parasitol., 84, 50, 1998. 33. Marsh, A.E. et al. Sequence analysis and polymerase chain reaction amplification of small subunit ribosomal DNA from Sarcocystis neurona. Am. J. Vet. Res., 57, 975, 1996. 34. Kaufmann, H. et al. Discrimination of Neospora caninum from Toxoplasma gondii and other apicomplexan parasites by hybridization and PCR. Mol. Cell. Probes, 10, 289, 1996. 35. MacPherson, J.M. and Gajadhar, A.A. Ribosomal RNA sequences for the specific detection of Toxoplasma gondii by hybridization assay. Mol. Cell. Probes. 7, 97, 1993. 36. Holmdahl, O.J. et al. Oligonucleotide probes complementary to variable regions of 18S rRNA from Sarcocystis species. Mol. Cell. Probes, 7, 481, 1993. 37. Beltrami, E.M. et al. Risk and management of blood-borne infections in health care workers. Clin. Microbiol. Rev., 13, 385, 2000. 38. Update: universal precautions for prevention of transmission of human immunodeficiency virus, hepatitis B virus, and other bloodborne pathogens in health-care settings. MMWR, 37, 377, 1988. 39. Tungtrongchitr, A. et al. The potential usefulness of the modified Kato thick smear technique in the detection of intestinal sarcocystosis during field surveys. Southeast Asian J. Trop. Med. Public Health, 38, 232, 2007. 40. Garcia, L.S. and Current, W.L. Cryptosporidiosis: clinical features and diagnosis. Crit. Rev. Clin. Lab. Sci., 27, 439, 1989. 41. Pena, H.F., Ogassawara, S. and Sinhorini, I.L. Occurrence of cattle Sarcocystis species in raw kibbe from Arabian food establishments in the city of Sao Paulo, Brazil, and experimental transmission to humans. J. Parasitol., 87, 1459, 2001. 42. Fujino, T. et al. Detection of a small number of Cryptosporidium parvum oocysts by sugar flotation and sugar centrifugation methods. J. Vet. Med. Sci., 68, 1191, 2006. 43. Tanhauser, S.M. et al. Multiple DNA markers differentiate Sarcocystis neurona and Sarcocystis falcatula. J. Parasitol., 85, 221, 1999. 44. Dubey, J.P. et al. Prevalence of viable Toxoplasma gondii in beef, chicken, and pork from retail meat stores in the United States: risk assessment to consumers. J. Parasitol., 91, 1082, 2005. 45. Dubey, J.P. and Beattie, C.P. Toxoplasmosis of animals and man, pp. 1–213. CRC Press Inc., Boca Raton, FL, 1988. 46. Schares, G. et al. Oocysts of Neospora caninum, Hammondia heydorni, Toxoplasma gondii and Hammondia hammondi in faeces collected from dogs in Germany. Int. J. Parasitol., 35, 1525, 2005. 47. Schares, G. et al. Occurrence of Toxoplasma gondii and Hammondia hammondi oocysts in the faeces of cats from Germany and other European countries. Vet. Parasitol., 152, 34, 2008. 48. Hill, D.E. et al. Specific detection of Neospora caninum oocysts in fecal samples from experimentally-infected dogs using the polymerase chain reaction. J. Parasitol., 87, 395, 2001. 49. Zhao, X., Duszynski, D.W. and Loker, E.S. A simple method of DNA extraction for Eimeria species. J. Microbiol. Methods, 44, 131, 2001. 50. Ajzenberg, D. et al. Genetic diversity, clonality and sexuality in Toxoplasma gondii. Int. J. Parasitol., 34, 1185, 2004. 51. Horn, K., Ono, K. and Heydorn, A.O. In vitro excystation and cryoreservation of ovine and caprine Sarcocystis species. J. Protozool. Res., 1, 3, 1991. 52. Dubey, J.P. et al. Prevalence of Sarcocystis neurona sporocysts in opossums (Didelphis virginiana) from rural Mississippi. Vet. Parasitol., 95, 283, 2001.

738 53. Elsheikha, H.M. et al. Generally applicable methods to purify intracellular coccidia from cell cultures and to quantify purification efficacy using quantitative PCR. Vet. Parasitol., 135, 223, 2006. 54. Yang, Z.Q. et al. Analysis of the 18S rRNA genes of Sarcocystis species suggests that the morphologically similar organisms from cattle and water buffalo should be considered the same species. Mol. Biochem. Parasitol., 115, 283, 2001. 55. Morrison, D.A. et al. The current status of the small subunit rRNA phylogeny of the coccidia (Sporozoa). Int. J. Parasitol., 34, 501, 2004. 56. Jenkins, M.C. et al. The relationship of Hammondia hammondi and Sarcocystis mucosa to other heteroxenous cystforming coccidia as inferred by phylogenetic analysis of the 18S SSU ribosomal DNA sequence. Parasitology, 119, 135, 1999. 57. Ellis, T.J. et al. Phylogenetic relationships between Toxoplasma and Sarcocystis deduced from a comparison of 18S rDNA sequences. Parasitology, 110, 521, 1995. 58. Ellis, J.T. et al. Molecular phylogeny of Besnoitia and the genetic relationships among Besnoitia of cattle, wildebeest and goats. Protist, 151, 329, 2000. 59. Ellis, J. and Morrison, D. Effects of sequence alignment on the phylogeny of Sarcocystis deduced from 18S rDNA sequences. Parasitol. Res., 81, 696, 1995. 60. Carreno, R.A. et al. Phylogenetic analysis of coccidia based on 18S rDNA sequence comparison indicates that Isospora is most closely related to Toxoplasma and Neospora. J. Eukaryot. Microbiol., 45, 184, 1998. 61. Barta, J.R. et al. Phylogenetic relationships among eight Eimeria species infecting domestic fowl inferred using complete small subunit ribosomal DNA sequences. J. Parasitol., 83, 262, 1997. 62. Barta, J.R. et al. Molecular phylogeny of the other tissue ­coccidia: Lankesterella and Caryospora. J. Parasitol., 87, 121, 2001. 63. Franzen, C. et al. Taxonomic position of the human intestinal protozoan parasite Isospora belli as based on ribosomal RNA sequences. Parasitol. Res., 86, 669, 2000. 64. Dame, J.B. et al. Sarcocystis falcatula from passerine and psittacine birds: synonymy with Sarcocystis neurona, agent of equine protozoal myeloencephalitis. J. Parasitol., 81, 930, 1995. 65. Cutler, T.J. et al. Are Sarcocystis neurona and Sarcocystis falcatula synonymous? A horse infection challenge. J. Parasitol., 85, 301, 1999. 66. Rosenthal, B.M., Lindsay, D.S. and Dubey, J.P. Relationships among Sarcocystis species transmitted by New World opossums (Didelphis spp.). Vet. Parasitol., 95, 133, 2001. 67. Sundar, N. et al. Modest genetic differentiation among North American populations of Sarcocystis neurona may reflect expansion in its geographic range. Vet. Parasitol., 152, 8, 2008. 68. Asmundsson, I.M., Dubey, J.P. and Rosenthal, B.M. A genetically diverse but distinct North American population of Sarcocystis neurona includes an overrepresented clone described by 12 microsatellite alleles. Infect. Genet. Evol., 6, 352, 2006. 69. Caraguel, C.G. et al. Microheterogeneity and coevolution: an examination of rDNA sequence characteristics in Neoparamoeba pemaquidensis and its prokinetoplastid endosymbiont. J. Eukaryot. Microbiol., 54, 418, 2007. 70. Chae, J. S. et al. Two Theileria cervi SSU RRNA gene sequence types found in isolates from white-tailed deer and elk in North America. J. Wildl. Dis., 35, 458, 1999.

Molecular Detection of Foodborne Pathogens 71. Reddy, G.R. et al. Sequence microheterogeneity of the three small subunit ribosomal RNA genes of Babesia bigemina: expression in erythrocyte culture. Nucleic Acids Res., 19, 3641, 1991. 72. Dubey, J.P. et al. Biological and molecular characterization of Besnoitia akodoni n.sp. (Protozoa: Apicomplexa) from the rodent Akodon montensis in Brazil. Parassitologia, 45, 61, 2003. 73. Dubey, J.P. et al. Besnoitia oryctofelisi n. sp. (Protozoa: Apicomplexa) from domestic rabbits. Parasitology, 126, 521, 2003. 74. Yang, Z.Q. et al. Identification of Sarcocystis hominis-like (Protozoa: Sarcocystidae) cyst in water buffalo (Bubalus bubalis) based on 18S rRNA gene sequences. J. Parasitol., 87, 934, 2001. 75. Li, Q.Q. et al. A PCR-based RFLP analysis of Sarcocystis cruzi (Protozoa: Sarcocystidae) in Yunnan Province, PR China, reveals the water buffalo (Bubalus bubalis) as a natural intermediate host. J. Parasitol., 88, 1259, 2002. 76. Yang, Z. Q. et al. Characterization of Sarcocystis species in domestic animals using a PCR-RFLP analysis of variation in the 18S rRNA gene: a cost-effective and simple technique for routine species identification. Exp. Parasitol., 102, 212, 2002. 77. Black, W.C. and Vontas, J.G. Affordable assays for genotyping single nucleotide polymorphisms in insects. Insect Mol. Biol., 16, 377, 2007. 78. Fakhrai-Rad, H., Pourmand, N. and Ronaghi, M. Pyrosequencing: an accurate detection platform for single nucleotide polymorphisms. Hum. Mutat., 19, 479, 2002. 79. Gibson, N.J. The use of real-time PCR methods in DNA sequence variation analysis. Clin. Chim. Acta, 363, 32, 2006. 80. Kim, S. and Misra, A. SNP genotyping: technologies and biomedical applications. Annu. Rev. Biomed. Eng., 9, 289, 2007. 81. Barta, J.R. and Thompson, R.C.A. What is Cryptosporidium? Reappraising its biology and phylogenetic affinities. Trends Parasitol., 22, 463, 2006. 82. Wolfe, N.D., Dunavan, C.P. and Diamond, J. Origins of major human infectious diseases. Nature, 447, 279, 2007. 83. Yang, Z.-Q. et al. A taxonomic re-appraisal of Sarcocystis nesbitti (Protozoa: Sarcocystidae) from the monkey Macaca fascicularis in Yunnan, PR China. Parasit. Int., 54, 75, 2005. 84. Stolte, M., Odening, K. and Bockhardt, I. Antelopes (Bovidae) kept in European zoological gardens as intermediate hosts of Sarcocystis species. Parassitologia, 38, 565, 1996. 85. Odening, K., Stolte, M. and Bockhardt, I. On the diagnostics of Sarcocystis in cattle: sarcocysts of a species unusual for Bos taurus in a dwarf zebu. Vet. Parasitol., 66, 19, 1996. 86. Hendricks, L.D. Host range characteristics of the primate coccidian, Isospora arctopitheci Rodhain 1933 (Protozoa: Eimeriidae). J. Parasitol., 63, 32, 1977. 87. Dubey, J.P. et al. A review of Sarcocystis neurona and equine protozoal myeloencephalitis (EPM). Vet. Parasitol., 95, 89, 2001. 88. Yabsley, M.J. et al. Seroprevalence of Toxoplasma gondii, Sarcocystis neurona, and Encephalitozoon cuniculi in three species of lemurs from St. Catherines Island, GA, USA. Vet. Parasitol., 144, 28, 2007. 89. Mugridge, N.B. et al. Phylogenetic relationships of the genus Frenkelia: a review of its history and new knowledge gained from comparison of large subunit ribosomal ribonucleic acid gene sequences. Int. J. Parasitol., 29, 957, 1999.

Sarcocystis 90. Modry, D., Votypka, J. and Svobodova, M. Note on the taxonomy of Frenkelia microti (Findlay & Middleton, 1934) (Apicomplexa: Sarcocystidae). Syst. Parasitol., 58, 185, 2004. 91. Slapeta, J.R. et al. Evolutionary relationships among cystforming coccidia Sarcocystis spp. (Alveolata: Apicomplexa: Coccidea) in endemic African tree vipers and perspective for evolution of heteroxenous life cycle. Mol. Phylogenet. Evol., 27, 464, 2003. 92. Dolezel, D. et al. Phylogenetic analysis of Sarcocystis spp. of mammals and reptiles supports the coevolution of

739 Sarcocystis spp. with their final hosts. Int. J. Parasitol., 29, 795, 1999. 93. Dubey, J.P. et al. Morphologic and genetic characterization of Sarcocystis sp. from the African grey parrot, Psittacus erithacus, from Costa Rica. Acta Parasitol., 51, 161, 2006. 94. Dubey, J.P. et al. Molecular phylogeny implicates New World opossums (Didelphidae) as the definitive hosts of Sarcocystis ramphastosi, a parasite of the keel-billed toucan (Ramphastos sulfuratus). Acta Protozool., 47, 55, 2008.

53 Toxoplasma Chunlei Su

University of Tennessee

J.P. Dubey

United States Department of Agriculture

Contents 53.1 Introduction.....................................................................................................................................................................741 53.1.1 Taxonomy and Biology .....................................................................................................................................741 53.1.2 Epidemiology and Pathogenesis....................................................................................................................... 743 53.1.3 Current Laboratory Diagnosis.......................................................................................................................... 743 53.1.3.1 Serological Tests.............................................................................................................................. 743 53.1.3.2 Bioassays in Mice and Cats............................................................................................................. 744 53.1.3.3 Molecular Techniques...................................................................................................................... 745 53.2 Methods.......................................................................................................................................................................... 746 53.2.1 Reagents and Equipment................................................................................................................................... 746 53.2.2 Sample Preparation........................................................................................................................................... 746 53.2.2.1 Extract T. gondii DNA from Animal Tissues (Muscle, Spleen, Brain, etc.)................................... 746 53.2.2.2 Purify T. gondii Oocysts from Water Samples and Extract DNA................................................... 747 53.2.3 Detection Procedures........................................................................................................................................ 747 53.2.3.1 Detection by nPCR.......................................................................................................................... 747 53.2.3.2 Detection by Real-Time PCR.......................................................................................................... 748 53.2.3.3 Nested Multilocus PCR-RFLP Genotyping.................................................................................... 749 53.3 Conclusion and Future Perspectives............................................................................................................................... 750 References...................................................................................................................................................................................751

53.1  Introduction 53.1.1 Taxonomy and Biology “The phylum Apicomplexa covers over 4600 species of ­intracellular protozoan parasites. Several genera in this phylum (e.g., Plasmodium, Crypotosporidium, Sarcocystis, Eimeria, Neospora and Toxoplasma) are of medical and veterinary importance.1 Being the only species in the genus Toxoplasma, T. gondii infects nearly all warm-blooded vertebrates including mammals and birds.2–4” The name Toxoplasma (toxon = arc, plasma = form, Greek) was derived from its crescent shape. Toxoplasma was first discovered in the desert rodent Ctenodactylus gundi by Charles Nicolle and Louis Manceaux at the Institut Pasteur in Tunis in 1908.5 At about the same time, Alfonso Splendore independently described Toxoplasma in a laboratory rabbit in Sao Paulo.6 Initially the parasite was mistakenly identified to be Leishmania by both groups, but soon it was realized to be a new species and named Toxoplasma gondii based on its morphology and host.7 T. gondii has a complicated life cycle that includes sexual and asexual replications8–10,16 (Figure 53.1). Cats are the only

known definitive hosts in which sexual replication of parasite occurs. Warm-blooded vertebrates can serve as intermediate hosts in which asexual replication takes place.2,4 The resistant stage of the parasite, the oocysts are shed by cats into the environment where oocysts are ingested by intermediate hosts. After several days of replication as rapidly growing tachyzoites during acute infection, the parasites switch to slow-growing bradyzoites, which form tissue cysts and reside in host cells for the life of the host. In geographical areas of the world that lack cats, T. gondii infection is rare or absent.11–14 In addition, prevention of oocysts shedding in cats by vaccination can drastically reduce prevalence of T. gondii in rodent population on farms.14 All these data indicate that cats play a pivotal role in perpetuating and sustaining the life cycle of T. gondii. There are three infectious stages of T. gondii: tachyzoites, bradyzoites in tissue cysts and sporozoites in oocysts (Figure 53.2). The tachyzoites (tachos = speed in Greek) are the fast dividing forms that multiply rapidly in nucleated host cells. They are often in crescent shape and about 2 × 6 µm in size. Their anterior ends are pointed and the posteriors are round. Tachyzoites enter host cells by active penetration of host 741

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Molecular Detection of Foodborne Pathogens

Definitive host (cat)

Unsporulated oocysts passed in feces

Tissue cysts ingested by cat Tissue cysts in intermediate hosts Tissue cysts ingested in infected uncooked meat

Tachyzoites transmitted through placenta

Oocysts in feed, water, or soil

Contaminated food and water Intermediate hosts

Ingested by intermediate host Sporulated oocysts

Infected fetus

Figure 53.1  Life cycle of Toxoplasma gondii. (From Dubey, J.P. et al., Clin. Microbiol. Rev., 11, 267, 1998.) (a)

(b)

10µm

(d)

10µm

(c)

(e)

2µm

Figure 53.2  Oocysts, tachyzoites, and tissue cyst of Toxoplasma gondii. (a) Unsporulated oocyst. (b) Sporulated oocyst with two sporocysts. Four sporozoites are visible in one of the sporocysts (arrows). (c) Transmission electron micrograph of a sporulated oocyst, with a thin oocyst wall (large arrow), two sporocysts (arrowheads), and sporozoites (small arrows). (d) tachyzoites in parasitophorous vacuole in fibroblast cells (arrow). (e) mouse brain tissue cyst (arrow) containing hundreds of bradyzoites. (From Dubey, J.P. et al., Clin. Microbiol. Rev., 11, 267, 1998.)

cell membrane and become surrounded by a parasitophorous vacuole (PV) that is resistant to digestion by host cell defense mechanism.15 In PV, tachyzoite replicates asexually by repeated binary division (endodyogeny) until the host cell lysis. After an unknown number of rapid replications, the

tachyzoite switches to a slow growing phase and give rises to bradyzoite.2 The bradyzoite (brady = slow in Greek) is similar to tachyzoite but multiply slowly. Bradyzoite is slender and measuring 7 × 1.5 µm in size, it has the nucleus situated

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Toxoplasma

toward the posterior end whereas the nucleus in tachyzoite is more central.16 Bradyzoites in tissue cysts are the “resting” stage of the parasite. Tissue cysts grow slowly and vary in size from 5 to 200 µm and contain a few to more than a thousand bradyzoites.16 Though tissue cysts can develop in all tissues, they are more prevalent in muscular and neural tissues. When ingested by cats, the wall of tissue cyst is digested by gastric enzymes in the stomach and small intestine, and the bradyzoites are released. The freed bradyzoites undergo two different ways of replication and dissemination. Some of these bradyzoites penetrate the lamina propria of the small intestine and replicate as tachyzoites, which are disseminated to other tissues. Some other bradyzoites penetrate epithelial cells of the small intestine, undergo numerous generation of asexual replication. These parasites are structurally distinct from tachyzoites and these stages are designated as types A–E. Following the asexual development of type A–E, the parasites differentiate to form male and female gametes. The male gamete has two flagella and can swim to and fuse with a female gamete to form oocyst, which is released into intestinal lumen. It may take 3–10 days for a tissue cyst infected cat to produce and release oocysts.16 Oocysts are spherical in shape, 10 × 12 μm in size. Newly released oocysts in cat feces are unsporulated and noninfectious. Within 1–5 days after excretion in cat feces, the oocysts sporulate to produce eight sporozoites that are highly infectious to animals and humans.2 Cats can shed millions of oocysts within 1–2 weeks.2,17,18 Oocysts can survive for months to years under moderate environmental condition and their resistance to destruction assures potential widespread contamination of food and water supplies.3,19

53.1.2 Epidemiology and Pathogenesis Toxoplasma gondii is one of the most widely spread protozoan parasites. It can virtually infect all warm-blooded mammals and birds. About one-third of the human population are chronically infected.3,4 T. gondii infection may occur by one of the four routes: (i) oral ingestion of infectious oocysts from contaminated food or water, (ii) oral ingestion of tissue cysts contained in raw or undercooked meat, (iii) transplacental transmission of tachyzoites from mother to developing fetus, and (iv) transmission of tachyzoites or bradyzoites in tissue cysts through organ transplantation.3,4,20 Primary infections in healthy adults are mostly asymptomatic but in some patients clinical toxoplasmosis can occur. Primary infection in healthy women during pregnancy may cause severe congenital toxoplasmosis. Frequency and severity of congenital transmission vary according to the stage of gestation when the mother became infected. Infection during early pregnancy may result in severe disease and even death of the fetus. Late maternal infection often results in asymptomatic newborns, however, these babies may later develop chorioretinitis or delayed growth and leaning disabilities.20,21 In immunocompromised patients such as organ transplant recipients and patients with AIDS, encysted bradyzoites may be reactivated to become fast-growing tachyzoites that

can cause life-threatening encephalitis.21 In some patients, severe disseminated toxoplasmosis occurs in immunocompetent human patients, perhaps related to the strain of T. gondii.22–24

53.1.3 Current Laboratory Diagnosis Clinical symptoms of T. gondii infection are nonspecific and unreliable for diagnosis. Due to the lack of a reliable method in diagnosis, the high prevalence of T. gondii in humans and animals was not recognized in the first 40 years since the discovery of this parasite. With the development of a sensitive and reliable serum test in 1948,25 knowledge of T. gondii epidemiology expanded rapidly. Conventional diagnosis of T. gondii infection includes serologic tests, bioassay, or combination of the two methods. 53.1.3.1 Serological Tests A broad range of serologic tests have been developed for the detection of antibodies to T. gondii in human and animals.2,26 These tests vary in specificity and sensitivity. A few frequently used methods in detection of T. gondii infection in humans and animals are briefly discussed below. Sabin–Feldman dye test (DT). The DT test was developed by Albert Sabin and Harry Feldman in 1948.25 It is highly sensitive and specific and still is considered as the “gold standard” for diagnosis of T. gondii infection in humans. The DT test measures complement fixing antibodies. In this test, live tachyzoites are incubated with test serum in the presence of complement from human serum. After incubation, a dye, methylene blue is added. In the absence of T. gondii specific antibody, tachyzoites are stained with methylene blue. However, in the presence of specific antibody, complement-mediated cytolysis of tachyzoite occurs and there is no staining of cytoplasm. Though highly sensitive and specific in diagnosis of human Toxoplasma infection, the DT test is expensive and potentially unsafe because of using live tachyzoites. Indirect fluorescent antibody test (IFA). In the IFA test, inactivated tachyzoites are incubated with sample serum and then fluorescent-labeled antispecies IgG is added. Detection of T. gondii specific IgG is achieved using fluorescent microscope. The disadvantages of IFA are the need of fluorescent microscope and species specific labeling of antispecies IgG.2 Indirect hemagglutination test (IHA). This test was developed by Jacobs and Lunde in 1957.27 Soluble antigens are prepared from tachyzoites and coated on tanned red blood cells (RBCs). Then the sample serum is mixed with the RBCs. In the presence of antibodies to T. gondii antigens, the RBCs will agglutinate and give positive readout. The IHA test is simple but can be highly variable with lower specificity comparing to DT test.2 Enzyme-linked immunoabsorbent assay (ELISA). In the ELISA test, soluble antigen is coated to microtiter plates and sample serum is added to form an antigen–antibody complex (if specific antibodies are present). A secondary

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enzyme-linked antibody specific to the host species is added to detect antigen–antibody complex. This test requires an ELISA reader and also enzyme conjugation to the secondary antibodies. Numerous modifications of ELISA have been reported to enhance specificity and simplify the protocol of the conventional ELISA.2 The ELISA tests are widely used in the diagnosis of human congenital toxoplasmosis.26 Modified direct agglutination test (MAT). Dubey and Desmonts28 modified the direct agglutination test that was developed by Desmonts and Remington.29 In the MAT test, antigen (formalin killed tachyzoites) is mixed with 2-mercaptoethanol and then added to serum samples that are placed into microtiter plates. The 2-mercaptoethanol destroys natural IgM antibodies that cause false positive of agglutination test. This significantly improves the test specificity. The sensitivity and specificity of MAT has been validated by comparing serologic data and isolation of the parasite from naturally and experimentally infected pigs.30,31 Due to its ease of use and high specificity, the MAT test has been the preferred method for the diagnosis of Toxoplasma infection in animals. 53.1.3.2 Bioassays in Mice and Cats Serological tests make diagnosis of T. gondii infection indirectly by detecting antibodies. These tests may not be able to detect T. gondii infections in newborns when their immune systems are not mature to produce antibodies. The alternative diagnosis is to isolate the parasite from host tissue samples. This can be achieved by bioassays in mice or cats. Both mice and cats are natural hosts of T. gondii and they can serve as a biological incubator to produce a large number of parasites to ease the identification of infection. The benefit of bioassay is that the parasite can be expanded and cryopreserved for future studies. Bioassay in mice. For the isolation of T. gondii from body fluids, (blood, cerebrospinal fluid, aqueous humor, amniotic fluid, etc.), the samples can be centrifuged at 1000 × g for 10 min, the pellet (may be invisible) is resuspended in phosphate buffered saline (PBS), and up to 0.5 ml of suspension can be inoculated to mice by intraperitoneal (IP) or subcutaneous injection. After IP inoculation, parasite tissue burden may peak between days 6 and 10 post IP. Tachyzoites may be found in the peritoneal fluid, or in the mesenteric lymph nodes and lungs. To collect and expand parasites in cell culture, euthanize mice on day 7 and inject 5 ml PBS into the peritoneal cavity, massage mouse abdomen a few times and collect peritoneal lavage. Centrifuge peritoneal lavage at 1000 × g for 10 min and resuspend the pellet in 0.5 ml of PBS, then transfer the parasites into a T25 flask cell culture of human foreskin fibroblast (HFF) monolayer. It generally takes about 2–3 weeks for the parasites to expand in the cell culture. These parasites can be cryopreserved in liquid nitrogen for long term storage. Alternatively, mouse homogenate can be cryopreserved for long term storage. Parasites collected from mice can be digested by proteinase K and genotyped by PCR-restriction fragment length polymorphism (PCR-RFLP) method, which will be discussed in later sections.

Molecular Detection of Foodborne Pathogens

Animal tissues, often muscular in nature such as tongue, heart, and diaphragm, can be used to isolate T. gondii in mice. As the number of T. gondii in tissue cysts of chronically infected animals is normally low, it is necessary to digest these tissues by enzyme pepsin to release parasites and concentrate before infecting mice. Dubey modified a previously established pepsin digestion protocol32 and improved sensitivity in isolating T. gondii from a variety of animal tissues.33 In brief, 50 g of muscle in a blender are ground briefly at low speed without saline. Then 125 ml of saline is added and blended briefly at top speed to make tissue homogenate. The blender is rinsed with 125 ml saline and the washings added to the tissue homogenate. Tissue homogenate is warmed to 37ºC, then mixed with 250 ml freshly prepared, pre-warmed (37ºC) acid pepsin solution (pepsin, 2.6 g; NaCl, 5 g; HCl, 7 ml and add distilled water to make 500 ml, pH ~1.1–1.2). The mixture is incubated at 37ºC for 60 min with shaking. Filter the homogenate through two layers of gauze, divide into two bottles, each with 250 ml of homogenate and centrifuge at 1200 × g for 10 min. The supernatant is discarded, and the pellet is resuspended in 20 ml PBS (pH 7.2), neutralized with 1.2% sodium bicarbonate (pH 8.3). After mixing, centrifuge at 1200 × g for 10 min. Discard the supernatant, then add 10 ml PBS that contains 1000 U/ml penicillin and 100 μg/ ml of streptomycin. Inoculate 1 ml of tissue homogenate subcutaneously or 0.5 ml IP into each of five to ten outbred mice. Between 6 and 10 days post infection, T. gondii can be isolated as described above. To prevent contamination, all equipment used in the above protocol are disinfected with hot water (70–100ºC), and all disposable biological wastes are autoclaved. Bioassay in cats. Often the number of tissue cysts in animals may be too low to be detectable by bioassay in mice. To increase the possibility of isolating the parasites, cats are usually used instead because they can consume much larger volume of tissues (ten times or more).2 As the end product of procedure is oocyst, and the sporulated oocyst is highly infectious to human, the protocol and equipment used for this purpose must be certified by the institutional biosafety requirements. After termination of the experiment, cats are euthanized and cat cages are autoclaved before cleaning. Any waste generated in the process of oocyst collection are autoclaved before disposal. To bioassay in cats, 500 g or more meat samples can be fed to specific pathogen free cats. Infected cats can shed millions of oocysts between days 3 and 14 post infection. During this period of time, all feces should be collected. The following is the outline for isolating oocysts from cat feces.2 Collect feces, add enough water and soak until feces have softened. Emulsify the feces and add 10 volumes of sucrose solution (sucrose 53 g, water 100 ml, liquid phenol 0.8 ml). Filter feces through two layers of gauze and pour the filtered feces into a 15 ml centrifuge tube. Centrifuge at 400 × g for 10 min, collect 0.5 ml from the very top of the tube and mix with 4.5 ml of 2% sulfuric acid in a 30 ml bottle. Cap the bottle and incubate at room temperature for 3–7 days to sporulate oocysts. After sporulation, transfer 20 μl

Toxoplasma

of oocysts to 500 μl PBS (pH 7.2) in a 1.5 ml centrifuge tube, mix well and centrifuge at 400 × g for 5 min. Discard supernatant and resuspend in 500 μl PBS. IP inject five mice each with 100 μl oocysts. Between day 7 and 10 post infection, collect peritoneal lavage from mice and inoculate tachyzoites to T25 HFF cells to expand the parasites for cryopreservation, or prepare cell lysate for PCR-RFLP genotyping. The diagnosis of toxoplasmosis traditionally depends on serology and the isolation of parasites in biological samples. Serological techniques sometimes are unreliable. Mouse inoculation is considered the standard method, because of its high sensitivity and specificity. However, this technique is time consuming, which is of concern for prompt diagnosis, and also the need for facilities to inoculate the animals. To overcome these limitations, PCR based molecular techniques have been developed over the past two decades. Today these methods are widely used in identification of T. gondii infections in humans and animals. A wide range of biological samples have been tested including amniotic fluid, cerebrospinal fluid, aqueous humor, serum, and blood and a variety of tissues. 53.1.3.3  Molecular Techniques Direct detection of parasite-specific DNA can be achieved by PCR. In general, PCR methods have proved to be simple, rapid, sensitive, reproducible, and cost-effective, which can be applied to a variety of clinical samples.34–37 PubMed search using key words “Toxoplasama” and “polymerase chain reaction (PCR)” between the years 2000 and 2008 retrieved more than 150 reference papers reporting PCR based detection and diagnosis of T. gondii in humans and animals, indicating its wide use in the field. However, up to date, there is no consensus on a standard set of DNA targets for PCR-based detection. Among many different markers, the 35-copy B1 gene and the 300-copy 529-bp element are most frequently used. The 110-copy internal transcribed spacer (ITS1) sequence was also successfully but less frequently applied for diagnosis. Molecular detection of T. gondii by the 35-copy B1 gene was first reported about two decades ago,38 and since then it has been modified and adapted in numerous laboratories and only a few are referenced here.35,36,39–46 Molecular methods can be divided into two categories. The first category focuses on high detection sensitivity. Its goal is to confirm the existence of the pathogen of interest. To reach this goal, highly repetitive DNA sequences, short replication targets and nested PCR are combined. An alternative is to use highly repetitive sequences, short replication targets and real-time PCR. However, these methods can not provide any information regarding strain type of the samples. The second category of molecular technique focuses on multilocus genotyping of single copy genes, with its goal of identifying each positive sample with high resolution. This is of importance for epidemiology studies that need to trace the path of disease transmission and identify the source of contamination. The disadvantage of using single-copy gene target is that the sensitivity is compromised when comparing

745

to the highly repetitive sequences. However, this can be compensated to some extent by nested PCR. Nested PCR (nPCR) of repetitive DNA sequences. The nPCR was applied for B1 and ITS1 sequences,35,36,42,44,46–48 both showed a similar sensitivity and can detect as low as one parasite/PCR reaction.37,47,48 The 529-bp repeat element was identified from Toxoplasma genome in 2000 by Homan et al., and this marker is 10–100 times more sensitive than the B1 marker.37,43,49 Because of its high sensitivity, the 529-bp element becomes a preferred marker for the detection of T. gondii in human and animal tissue samples. Real-time PCR of repetitive DNA sequences. In recent years, advances in real-time PCR technology provided new approach in detecting pathogens in biological samples. The advantage of this method is that it not only detects but also quantifies the pathogens. Because of that, real-time PCR is gaining popularity in detection of T. gondii.34,35 This technique is quantitative, gives results over a range of six to seven orders of magnitude with low intra- and inter-assay variation.35,48,50 The detection threshold of real-time PCR in T. gondii is superior to nPCR assays.35 Quantification of T. gondii DNA by real-time PCR has been achieved from infected pig and mouse tissues48 and from clinical samples of whole blood and amniotic fluid.35,50,51 Real-time PCR is extremely useful when standard methods (usually microscopical or serological) for quantification are difficult or are less sensitive for detection of clinically relevant parasite burdens. The ability to quantify low-level parasite burdens allows the monitoring of the progression of infections under drug treatment and diagnosis of low-level infections or carrier states.35 Multilocus nested PCR-RFLP genotyping. Whereas both nested and real-time PCRs are highly sensitive and specific in the diagnosis of T. gondii in biological samples, they provide no information regarding the genotypes of the parasite, which is very important in the understanding of epidemiology in T. gondii. Previous studies on molecular epidemiology in T. gondii had shown that the parasite had a clonal population structure with the type I, II, and III lineages dominant in North America and Europe,52–56 whereas the parasites in South America are highly diverse with a few genotypes expanded into a wider range of geographical regions.57–59 These data indicate different population structures exist in different geographical regions. The consequences of infections with T. gondii may depend on the host genetic makeup and parasite genotypes. In humans, disease manifestations vary widely, ranging from asymptomatic to severe acute toxoplasmosis.21,23 Type II lineage is the predominant type causing human toxoplasmosis.53,56 Limited data suggest that there is association of disease presentations and parasite genotypes. For examples, type I or type I-like atypical strains are more likely to be involved in severe toxoplasmic retinochoroiditis or AIDS patients60,61 and atypical strains often cause severe acute disseminated toxoplasmosis in immunocompetent patients.23,24 This challenges the currently accepted practice in that no medical treatment is necessary when healthy individuals are exposed to T. gondii. Understanding the association between parasite genotypes

746

and diseases is essential in making appropriate decisions in treatment and control of toxoplasmosis. However, the prerequisite for this measure is correct identification of each T. gondii isolate by high resolution genotyping. There are over 200 genetic markers that can be used for genotyping in T. gondii.62,63 Most of these are PCR-RFLP markers.53,59,63–67 A small number of markers are microsatellites which have different copy number of di-nucleotide repeats.54-56,58,68–70 The high diversity of T. gondii strains world­wide requires a high resolution typing method for epidemiological studies. Such a method should be easy and simple to perform by ordinary laboratories without sophisticated equipments. To this end, a multilocus nested PCR-RFLP method (include four markers) was initially tested and showed that the sensitivity of the assay was less than ten parasites per PCR reaction when the parasites were spiked in normal human cerebrospinal fluid.61 Similar approach was taken to directly detect and genotype T. gondii strains in cerebrospinal fluid of human AIDS patients in Brazil.67 Based on these results, a multilocus nested PCR-RFLP typing method with ten genetic markers was developed, which has a higher resolution in identifying T. gondii isolates collected from different hosts in a variety of geographical regions.59,64,65 In this method, all ten markers are pre-amplified by multiplex PCR using external primers, then the pre-amplified PCR products are used to amplify each individual marker by nPCR. The advantage is that only a few microliter samples are needed. This method has been successfully applied to genotype T. gondii directly from a number of sea otter brain samples71 and from over 50 arctic fox brain samples collected in Svalbard islands near the arctic circle76. Taken together, these data prove that it is feasible to genotype T. gondii isolates from animal tissue samples by nested multilocus PCR-RFLP method. Application of this method to a broad range of samples will greatly facilitate our understanding of molecular epidemiology and population diversity of T. gondii in the near future.

53.2  Methods Based on the advances and our knowledge of molecular methods available for T. gondii, we suggest an integrated approach to detect and genetically characterize the parasite samples. This will include nested or real-time PCR of repetitive DNA sequences B1 and the 529-bp repeat elements, and multilocus nested PCR-RFLP genotyping of the samples.

53.2.1 Reagents and Equipment Molecular detection of T. gondii can be achieved by nPCR or real-time PCR. The nPCR can be performed using a regular thermal cycler (~$5000–$10000). The real-time PCR will need a more expensive thermal cycler (~$30000) which is able to detect fluorescent signals and quantify DNA during the runs of PCR reactions. In this section we will use Roche LightCycler as an example because available real-time PCR protocols for T. gondii were mostly developed based on this model.

Molecular Detection of Foodborne Pathogens

Reagents for nPCR and the detection of PCR products include DNA grade H2O, PCR buffer, dNTPs, PCR primers, hot start Taq DNA polymerase, DNA markers, DNA loading dye and ethidium bromide. The basic equipment needed are thermal cycler and gel electrophoresis equipment. For realtime PCR, additional reagents including dual fluorescence resonance energy transfer (FRET) probes specific to the target DNA sequence are needed. A real-time PCR machine such as the LightCycler is necessary. As PCR is sensitive to contamination, common practice should apply to reduce contamination to the minimum. These include separating the zones for PCR preparation, amplification, electrophoresis of final products and discarding of waste. Amplified PCR products should never be brought to the zone for PCR preparation. To monitor contamination, negative controls should be included with test samples at all stages.

53.2.2 Sample Preparation 53.2.2.1 Extract T. gondii DNA from Animal Tissues (Muscle, Spleen, Brain, etc.) Materials: QIAamp DNA Mini Kit (QIAamp Mini Spin Columns, 2-ml collection tubes, Buffer AL, Buffer ATL, Buffer AW1, Buffer AW2, Buffer AE, Proteinase K). Ethanol (96–100%) 1.5 ml, 2 ml microcentrifuge tubes Pipette tips with aerosol barrier Microcentrifuge (with rotor for 2 ml tubes) Vortexer Water bath or heating block at 56°C Procedure: (1) Cut up to 25 mg of tissue (up to 10 mg spleen) into small pieces. Place in a 1.5-ml microcentrifuge tube. (2) Add 180 µl Buffer ATL. (3) Add 20 μl proteinase K, mix by vortexing, and incubate at 56°C until the tissue is completely lysed. Vortex occasionally during incubation to disperse the sample. (4) Briefly centrifuge the 1.5-ml microcentrifuge tube to remove drops from the inside of the lid. (5) Add 200 μl Buffer AL to the sample, mix by pulsevortexing for 15 sec, and incubate at 70°C for 10 min. Briefly centrifuge the 1.5-ml microcentrifuge tube to remove drops from inside the lid. (6) Add 200 μl ethanol (96–100%) to the sample, and mix by pulse-vortexing for 15 sec. After mixing, briefly centrifuge the 1.5-ml microcentrifuge tube to remove drops from inside the lid. (7) Carefully apply the mixture to the 2-ml QIAamp Mini Spin Column. Centrifuge at 6000 × g for

747

Toxoplasma









1 min. Place the QIAamp Mini Spin Column in a clean 2-ml collection tube, and discard the tube containing the filtrate. (8) Add 500 μl Buffer AW1, centrifuge at 6000 × g for 1 min. Place the QIAamp Mini Spin Column in a clean 2-ml collection tube, and discard the collection tube containing the filtrate. (9) Add 500 μl Buffer AW2 and centrifuge at full speed (10000 × g) for 3 min. (10) Place the QIAamp Mini Spin Column in a new 2-ml collection tube and discard the old collection tube with the filtrate. Centrifuge at full speed for 1 min. (11) Place the QIAamp Mini Spin Column in a clean 1.5-ml microcentrifuge tube, and discard the collection tube containing the filtrate. Add 50 μl Buffer AE or distilled water to the QIAamp Mini Spin Column. Incubate at room temperature for 1 min, and then centrifuge at 6000 × g for 1 min. (12) Store DNA samples at –20°C before detection of T. gondii by nPCR, real-time PCR or multilocus nested PCR-RFLP genotyping.

Note: This protocol may be adapted to detect T. gondii in vegetables and fruits by washing these samples with 1 liter of PBS with 0.01% Tween 80, and then follow the above protocol in Section 53.2.2.2.

53.2.3  Detection Procedures In this section, molecular protocols for T. gondii detection and genotyping are outlined. They may be modified in individual laboratory to optimize the efficiency of PCR amplification. 53.2.3.1 Detection by nPCR Materials: FastStart Taq DNA polymerase Kit (Roche Applied Science) 2.5 mM dNTPs mix (containing 2.5 mM each dATP, dTTP, dCTP, and dGTP). Primers for nPCR of B1 and 529-bp repeat sequences are summarized in Table 53.1. Extracted DNA samples. Procedure:

53.2.2.2 Purify T. gondii Oocysts from Water Samples and Extract DNA This protocol is modified from Sroka et al.72 Materials: Sugar solution, specific gravity, 1.15 (sugar, 53 g, phenol 0.8 ml, H2O 100 ml) PBS with 0.01% Tween 80, pH 7.8 Distilled H2O 50 ml and 100 ml centrifuge tubes Microcentrifuge QIAamp DNA Mini Kit (as above) Procedure: (1) Collect 5 l of water sample, filter through cellulose filter with a pore diameter of 0.45 µm. (2) Wash the filter with 40 ml phosphate buffer (PBS, with 0.01% Tween 80, pH 7.8). (3) Discard filter and centrifuge at 1000 × g for 10 min. (4) Discard supernatant and resuspend the pellet in 20 ml of distilled water. (5) Transfer the pellet suspension to 30 ml sugar solution and centrifuge at 1000 × g for 10 min. (6) Collect 25 ml of the liquid from the surface to a new 100-ml centrifuge tube, add 75 ml PBS containing 0.01% Tween 80. (7) Centrifuge at 1000 × g for 10 min. Discard the supernatant. Resuspend pellet in 180 µl Buffer ATL from QIAamp DNA Mini Kit. (8) Follow step 3 from Section 53.2.2.1 to extract DNA.

(1) Thaw all PCR reagents (except DNA polymerase) and template DNA samples at room temperature for 20 min, tap each tube to mix it well, spin briefly. (2) Assemble the first PCR mix (number of reactions + 1) as following: H2O 10 × PCR buffer (Mg−) dNTPs (2.5 mM each) 25 mM MgCl2

26.2 µl 5.0 µl 4.0 µl 4.0 µl

External forward primer (50 µM)

0.30 µl

External reverse primer (50 µM)

0.30 µl

FastStart Taq (5 U/µl) Total

0.20 µl 40.0 µl

(3) Aliquot 40 µl of PCR mix to 0.2 ml PCR plate. Add 10 µl of extracted DNA sample. Spin briefly. (4) Heat samples at 95ºC for 5 min, then PCR 30 cycles of 94ºC for 30 sec, 55ºC for 1 min, 72ºC for 2 min. Soak at 4ºC. (5) Assemble nested PCR mix (number of reactions + 1) as following: H2O

31.2 µl

10 × PCR buffer (Mg–) dNTPs (2.5 mM each)

5.0 µl

25 mM MgCl2

4.0 µl

4.0 µl

Nested forward primer (50 µM)

0.30 µl

Nested reverse primer (50 µM)

0.30 µl

FastStart Taq (5 U/µl) Total

0.20 µl 45.0 µl

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Molecular Detection of Foodborne Pathogens

Table 53.1 The nPCR Primers of B1 Gene and 529-bp Element Markers (GenBank no.)

Primers

Primer Sequences

PCR (bp)

References

B1 (AF179871)

External

T8: ATGTGCCACCTCGCCTCTTGG T5: GCAATGCTTCTGCACAAAGTG T2: TGCATAGGTTGCAGTCACTG, T7: TAAAGCGTTCGTGGTCAACT Tox-8: GACGTCTGTGTCACGTAG ACCTAAG Tox-5: CTGCAGACACAGTGCATCTGGATT Tox-9: AGGAGAGATATCAGGACTGTAG, Tox-11: GCGTCGTCTCGTCTAGATCG

797

44,75

126

44,75

450

43,49

162

43,49

Internal 529-bp (AF487550)

External Internal

Table 53.2 Primers and Probes for Real-Time PCR of B1 Gene and The 529-bp Repeat Sequences Markers (GenBank no.) B1 (AF179871)

529-bp (AF487550)

Primer and Probe Sequences Forward: GGAGGACTGGCAACCTGGTGTCG Reverse: TTGTTTCACCCGGACCGTTTAGCAG Probe 1: CGGAAATAGAAAGCCATGAGGCACTCC-[FL] Probe 2: [Red 640]-ACGGGCGAGTAGCACCTGAGGAGAT-Ph Probe type: FRET Tox-9: AGGAGAGATATCAGGACTGTAG Tox-11: GCGTCGTCTCGTCTAGATCG Tox-HP-1: GAGTCGGAGAGGGAGAAGATGTT-[FL] Tox-HP-2: [Red 640]-CCGGCTTGGCTGCTTTTCCTG-Ph Probe type: FRET

PCR (bp)

References

126

73

162

43

[FL], fluorescein; [Red 640], LightCycler-Red 640-N-hydroxy-succinimide ester; [Ph], 3′-phosphate. FRET, fluorescence resonance energy transfer hybridization probes.

(6) Aliquot 45 µl PCR mix to 0.2 ml PCR plate. Add 5 µl of first PCR products. Spin briefly. (7) Heat samples at 95ºC for 5 min, then PCR 35 cycles of 94ºC for 30 sec, 55ºC for 1 min, 72ºC for 1 min. Soak at 4ºC. (8) To check nPCR products, take 5 µl of sample, mix with 2 µl of 5 × loading buffer and load to 2% agarose gel (containing 0.3 µg/ml ethidium bromide). Run gel in 1 × TBE buffer at 120 V for ~1 h. Visualize DNA bands under UV light. 53.2.3.2 Detection by Real-Time PCR This protocol is modified from Costa et al.73 Materials: LightCycler System (Roche Applied Science) LightCycler Capillaries Standard benchtop microcentrifuge containing a rotor for 2.0 ml reaction tubes PCR template (genomic DNA or cDNA) LightCycler Fast-Start Enzyme

LightCycler Fast-Start Reaction Mix, containing reaction buffer, dNTP mix (with dUTP instead of dTTP), and 10 mM MgCl2 MgCl2 stock solution, 25 mM H2O, PCR grade Mixed PCR primers (5 µM each) (Table 53.2) Mixed HybProbes (2.5 µM each) (Table 53.2) Procedure: (1) Thaw one vial of “Reaction Mix.” Briefly centrifuge one vial “Enzyme”. (2) Pipet 60 µl from “Reaction Mix” to “Enzyme” vial. Mix gently by pipetting up and down. (3) The new mix is 10 × LightCycler FastStart DNA Master HybProbe. (4) Thaw the solutions and, for maximal recovery of contents, briefly spin vials in a microcentrifuge before opening. (5) In a 1.5-ml reaction tube on ice, prepare the PCR mix for one 20 µl reaction by adding the following components in the order listed below:

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Toxoplasma

H2O, PCR-grade

8.0 µl

MgCl2 (25 mM)

1.0 µl

Primer mix (5 µM each)

2.0 µl

Mixed HybProbe (2.5 µM each) LightCycler FastStart DNA Master HybProbe

2.0 µl

Total

15 µl

2.0 µl

(6) Mix carefully by pipetting up and down. Pipet 15 µl PCR mix into each precooled LightCycler Capillary. Add 5 µl of the DNA sample. Seal each capillary with a stopper. (7) Place the adapters (containing the capillaries) into a standard benchtop microcentrifuge. Place the centrifuge adapters in a balanced arrangement within the centrifuge. Centrifuge at 700 × g for 5 sec (3000 rpm in a standard benchtop microcentrifuge). (8) Transfer the capillaries into the sample carousel of the LightCycler Instrument. (9) Pre-incubate the samples at 95°C for 10 min. Cycle the samples 50 cycles of: denaturation at 95°C for 10 sec; ramp rate, 20°C/sec, annealing at 60°C for 10 sec; ramp rate, 20°C/sec, and extension at 72°C for 15 sec; ramp rate, 20°C/sec. (10) Record and analyze data. 53.2.3.3  Multilocus nested PCR-RFLP Genotyping For those samples tested positive by nPCR or real-time PCR, they can be further identified by multilocus nested PCR-RFLP genotyping. Ten PCR-RFLP markers have been widely used in recent years in identifying T. gondii isolates from a variety of hosts and geographical regions.59,64,65 These markers are SAG1, SAG2, SAG3, BTUB, GRA6, c22-8, c29-2, L358, PK1, and Apico. For SAG2 gene, three sequence fragments including 5′-SA2, 3′-SAG2 and SAG2-new are analyzed. 5′-SA2 and 3′-SAG2 were frequently used and together they can distinguish the clonal type I, II, and III strains.74 The SAG2-new was developed recently which can identify additional allele not exists in the clonal type I, II, and III lineages.64 Materials: FastStart DNA polymerase kit (Roche Applied Science) dNTPs mix, 2.5 mM each External and internal primers each at 500 µM (Table 53.3) Thermocycler Procedure: (1) Mix 10 µl of 500 µM external forward primers, bring the final volume to 240 µl by adding H2O. The final concentration for each external forward primer is 21 µM 5′-SAG2 sequence. Prepare reverse primer mix by the same way. Note: 5′-SAG2 primers not needed as is covered by SAG2-new primers.

(2) Assemble multiplex PCR mix (number of reactions + 1) as following: 16.6 µl

H2O

2.5 µl

10 × PCR buffer (Mg−) dNTPs (2.5 mM each)

2.0 µl 2.0 µl

25 mM MgCl2 Mixed forward primers (21 µM each)

0.15 µl

Mixed reverse primers (21 µM each)

0.15 µl

FastStart Taq (5 U/ µl) Total

0.10 µl 23.5 µl

(3) Aliquot 23.5 µl of PCR mix to 0.2-ml PCR tubes. Add 1.5 µl of extracted DNA sample. Negative and positive controls are included for each batch of PCR. Spin briefly if necessary. (4) Heat samples at 95ºC for 4 min, then PCR 30 cycles of 94ºC for 30 sec, 55ºC for 1 min, 72ºC for 2 min. Soak at 4ºC. (5) Add 25 µl of H2O to dilute amplified PCR products. (6) Prepare nPCR mix (number of reactions + 1) as following: H2O

16.6 µl

10 × PCR buffer (Mg−) dNTPs (2.5 mM each)

2.5 µl

25 mM MgCl2

2.0 µl

Nested forward primer (50 µM)

0.15 µl

2.0 µl

Nested reverse primer (50 µM)

0.15 µl

FastStart Taq (5 U/ µl) Total

0.10 µl 23.5 µl

(7) Aliquot 23.5 µl of PCR mix to 0.2-ml microtubes. Add 1.5 µl of primary PCR products. Spin briefly if necessary. (8) Heat samples at 95ºC for 4 min, then PCR 35 cycles of 94ºC for 30 sec, 60ºC for 1 min, 72ºC for 1.5 min. Soak at 4ºC. (9) To check PCR products, take 5 µl of samples, mix with 2 µl of 5 × loading buffer and run 1.5% agarose gel with 0.3 μg/ml ethidium bromide in 1 × TBE buffer at 120 V for ~1.5 h. (10) Prepare digestion mix (number of reactions + 1): H2O

14.8 µl

10 × NEB buffer

1.8 µl

100 × BSA Restriction enzyme 1

0.2 µl

Restriction enzyme 2 (if apply) Total

0.1 µl 0.1 µl 17.0 µl

750

Molecular Detection of Foodborne Pathogens

Table 53.3 Summary of PCR Primers for Multilocus Nested PCR-RFLP Markers

Markers

External Primers (Multiplex PCR)*

Internal Primers (Nested PCR)

Nested PCR (bp)

Restriction Enzymes, NEB Buffers, Incubation Temperature and Time

Reference

SAG1

F: GTTCTAACCACGCACCCTGAG R: AAGAGTGGGAGGCTCTGTGA

F: CAATGTGCACCTGTAGGAAGC R: GTGGTTCTCCGTCGGTGTGAG

390

5′-SAG2

External primers not needed

242

3′-SAG2

F: TCTGTTCTCCGAAGTGACTCC R: TCAAAGCGTGCATTATCGC F: CAACTCTCACCATTCCACCC R: GCGCGTTGTTAGACAAGACA F: TCCAAAATGAGAGAAATCGT R: AAATTGAAATGACGGAAGAA

F: GAAATGTTTCAGGTTGCTGC R: GCAAGAGCGAACTTGAACAC F: ATTCTCATGCCTCCGCTTC R: AACGTTTCACGAAGGCACAC F: TCTTGTCGGGTGTTCACTCA R: CACAAGGAGACCGAGAAGGA F: GAGGTCATCTCGGACGAACA R: TTGTAGGAACACCCGGACGC

Sau96I + HaeII (double digest) NEB4, BSA, 37C 1 h MboI, NEB4, BSA, 37C 1 h

222

HhaI, NEB4, BSA, 37C 1 h

74

225

NciI, NEB4, BSA, 37C 1 h

60

411

63,64

F: ATTTGTGTTTCCGAGCAGGT R: GCACCTTCGCTTGTGGTT F: TGATGCATCCATGCGTTTAT R: CCTCCACTTCTTCGGTCTCA

F: TTTCCGAGCAGGTGACCT R: TCGCCGAAGAGTTGACATAG F: TCTCTCTACGTGGACGCC R: AGGTGCTTGGATATTCGC

344

BsiEI + TaqI (double digest), NEB4, BSA, 60C 1 h MseI, NEB2, BSA, 37C 1 h

521

BsmAI + MboII (double digest), NEB2, BSA, 37°C 30 min, 55°C 30min

63,64

c29-2

F: ACCCACTGAGCGAAAAGAAA R: AGGGTCTCTTGCGCATACAT

F: AGTTCTGCAGAGTGTCGC R: TGTCTAGGAAAGAGGCGC

446

HpyCH4IV + RsaI (double digest), NEB1, BSA, 37°C 1 h

63,64

L358

F: CTCTCGACTTCGCCTCTTC R: GCAATTTCCTCGAAGACAGG

F: AGGAGGCGTAGCGCAAGT R: CCCTCTGGCTGCAGTGCT

418

HaeIII + NlaIII (double digest), NEB4, BSA, 37°C 1 h

63,64

PK1

F: GAAAGCTGTCCACCCTGAAA R: AGAAAGCTCCGTGCAGTGAT

F: CGCAAAGGGAGACAATCAGT R: TCATCGCTGAATCTCATTGC

903

AvaI + RsaI (double digest), NEB4, BSA, 37°C 1 h

63,64

SAG2-new

F: GGAACGCGAACAATGAGTTT R: GCACTGTTGTCCAGGGTTTT

F: ACCCATCTGCGAAGAAAACG R: ATTTCGACCAGCGGGAGCAC

546

HinfI + TaqI, NEB3, BSA, 37°C 30 min, 65°C 30 min

63,64

Apico

F: TGGTTTTAACCCTAGATTGTGG F: TGCAAATTCTTGAATTCTCAGTT R: AAACGGAATTAATGAGATTTGAA R: GGGATTCGAACCCTTGATA

640

AflII + DdeI (double digest), NEB2, BSA, 37°C 1 h

64

SAG3 BTUB GRA6 c22-8

60 64,74

63,64

* F = forward primer; R = reverse primer.

(11) Aliquot 17 µl of enzyme mix to 0.5 ml tubes, add 3 µl PCR products, incubate for 1 h at the corresponding temperature. (12) Add 4 µl of 5 × loading dye, run 2.5% agarose gel in the presence of 0.3 µg/ml ethidium bromide in 1 × TBE buffer at 120 V for ~1.5 h. Note: for marker Apico, run 3% gel. Save gel pictures and record results. (13) Determine genotype results by comparing samples with the positive controls. To monitor contamination of PCR amplification, negative controls without DNA template added should be included in each batch of experiment. In addition, a set of positive controls that are prepared from cell culture are also included to monitor efficiency of PCR amplification and genotyping. The suggested positive controls include RH88, PTG, CTG, TgCgCa1 (a.k.a. COUGAR), MAS and TgCatBr5 and the genotyping data is summarized in Table 53.4. Examples of PCR-RFLP gel images for type I, II, and III strains RH88, PTG, and CTG are presented in Figure 53.3.64

53.3 Conclusion and Future Perspectives The techniques of molecular detection of T. gondii are well developed and widely used in clinical and basic research settings. This is evident by the large number of research papers published on this subject in the past several years. It is obvious that molecular methods become indispensable tools in the diagnosis and in the understanding of epidemiology of this parasite. However, at present there is no consensus on a standard protocol that all laboratories can follow, and therefore it is hard to compare data among different laboratories. To overcome this problem, a national quality assurance initiative in the molecular prenatal diagnosis of toxoplasmosis was established in France. Under this umbrella, panels of amniotic fluid containing varying concentrations of T. gondii were distributed to multiple laboratories for quality control in France. This approach provides the opportunity to improve laboratory practice and performance, and to increase communication among laboratories involved in making diagnosis.36 It is reasonable to expect that this type of communication and collaboration among different laboratories will be extended to the

751

Toxoplasma

Table 53.4 Summary of Multilocus PCR-RFLP Genotyping Results for T.gondii Positive Control Genetic Markers T. gondii Reference Strains

SAG1*

(5′ + 3′) SAG2†

SAG2-new‡

SAG3

BTUB

GRA6

RH88 (type I) PTG (type II) CTG (type III) TgCgCa1 (COUGAR) MAS TgCatBr5

I II or III II or III I u-1 I

I II III II I III

I II III II II III

I II III III III III

I II III II III III

I II III II III III

C22-8

C29-2

I II III II u-1 I

I II III u-1 I I

L358 I II III I I I

PK1

Apico

I II III u-2 III u-1

I II III I I I

* At SAG1 locus, type II and III are indistinguishable. SAG2 marker based on 5′- and 3′-ends of the gene sequence.74 ‡ A SAG2 marker based on the 5′-end of the gene sequence but different from 5′-SAG2.64 u-1 and u-2 are new alleles that are different from the clonal Type I, II, and III alleles.

†

c22-8 bp

M I II III

c29-2

L358

I II III

I II III

PK1

SAG2-new

I II III M M I II III

GRA6

SAG3

I II III

BTUB

I II III I

Apico

II III M M I II III

SAG1

5´-SAG2

3´-SAG2

I II III I II III I

II III

603 310 281 194 118 72

Figure 53.3  PCR-RFLP gel image. Reference type I, II, and III strains are RH88, PTG, and CTG, respectively. DNA from each marker is digested with corresponding restriction enzyme(s). All makers are resolved by running 2.5% gel, except Apico was resolved in 3% gel. M: DNA molecular marker phiX174/HaeIII digest.

whole Toxoplasma community (clinical and basic research) in the near future. At present, most molecular methods are focusing on detecting the presence of T. gondii in a variety of biological samples, but little is done to further characterize the parasites if present. This is due to the fact that all sensitive markers have multiple copies of the sequences, with B1 gene has 35 copies, ITS1 has 110 copies and the 529-bp element has over 300 repeats. In addition, in order to achieve maximum sensitivity, small-size PCR amplicons are favored, usually pushed to the limit with less than 200 bp. These markers normally have very limited polymorphisms and are not appropriate to differentiate parasite strains. The recent development of multilocus nested PCR-RFLP method makes it possible to genotype T. gondii isolates with high resolution. Combination of sensitive detection with high resolution genotyping will provide important information regarding epidemiology of T. gondii and relationship of disease manifestations with parasite genotypes. This in turn will improve the measures in control and treatment of T. gondii infections.

References





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9. Dubey, J.P., Miller, N.L., and Frenkel, J.K. The Toxoplasma gondii oocyst from cat feces. J. Exp. Med., 132, 636, 1970. 10. Hutchison, W. et al. Coccidian-like nature of Toxoplasma gondii. Brit. Med. J., 1, 142, 1970. 11. Wallace, G.D. Serologic and epidemiologic observations on toxoplasmosis on three pacific atolls. Am. J. Epidemiol., 90, 103, 1969. 12. Munday, B.L. Serological evidence of Toxoplasma infection in isolated groups of sheep. Res. Vet. Sci., 13, 100, 1972. 13. Dubey, J.P. et al. Low seroprevalence of Toxoplasma gondii in feral pigs from a remote island lacking cats. J. Parasitol., 83, 839, 1997. 14. Mateus-Pinilla, N.E. et al. A field trial of the effectiveness of a feline Toxoplasma gondii vaccine in reducing T. gondii exposure for swine. J. Parasitol., 85, 855, 1999. 15. Sibley, L.D. et al. Toxoplasma modifies macrophage phagosomes by secretion of a vesicular network rich in surface proteins. J. Cell Biol., 103, 867, 1986. 16. Dubey, J.P., Lindsay, D.S., and Speer, C.A. Structures of Toxoplasma gondii tachyzoites, bradyzoites and sporozoites and biology and development of tissue cysts. Clin. Microbiol. Rev., 11, 267, 1998. 17. Dubey, J.P. and Frenkel, J.F. Cyst-induced toxoplasmosis in cats. J. Protozool., 19, 155, 1972. 18. Dubey, J.P. Oocyst shedding by cats fed isolated bradyzoites and comparison of infectivity of bradyzoites of the VEG strain Toxoplasma gondii to cats and mice. J. Parasitol., 87, 215, 2001. 19. Dubey, J.P. Toxoplasma gondii oocyst survival under defined temperatures. J. Parasitol., 84, 862, 1998. 20. Jones, J.L. et al. Congenital toxoplasmosis: a review. Obstet. Gynecol. Surv., 56, 296, 2001. 21. Montoya, J.G. and Liesenfeld, O. Toxoplasmosis. Lancet 363, 1965-1976, 2004. 22. Carme, B. et al. Severe acquired toxoplasmosis in immunocompetent adult patients in French Guiana. J. Clin. Microbiol., 40, 4037, 2002. 23. Bossi, P. and Bricaire, F. Severe acute disseminated toxoplasmosis. Lancet, 364 (9434), 579, 2004. 24. Demar, M. et al. Fatal outbreak of human toxoplasmosis along the Maroni River: epidemiological, clinical, and parasitological aspects. Clin. Infect. Dis., 45, e88, 2007. 25. Sabin, A.B. and Feldman, H.A. Dyes as microchemical indicators of a new immunity phenomenon affecting a protzoan parasite (toxoplasma). Science, 108, 660, 1948. 26. Remington, J.S. et al. Toxoplasmosis. In Infectious Diseases of the Fetus and Newborn Infant, 6th ed. Remington, J.S. and Klein, J.O. (eds). W. B. Saunders Company, Philadelphia, PA, 2006, pp. 947–1091. 27. Jacobs, L. and Lunde, M.N. A hemagglutination test for toxoplasmosis. J. Parasitol., 43, 308, 1957. 28. Dubey, J.P. and Desmonts, G. Serological responses of equids fed Toxoplasma gondii oocysts. Equine Vet. J., 19, 337, 1987. 29. Desmonts, G. and Remington, J.S. Direct agglutination test for diagnosis of Toxoplasma infection: method for increasing sensitivity and specificity. J. Clin. Microbiol., 11, 562, 1980. 30. Dubey, J.P. et al. Sensitivity and specificity of various serologic tests for detection of Toxoplasma gondii infection in naturally infected sows. Am. J. Vet. Res., 56, 1030, 1995. 31. Dubey, J.P., Andrews, C. D., Thulliez, P., Lind, P., and Kwok, O.C.H. Long-term humoral antibody responses by various serologic tests in pigs orally inoculated with oocysts of four strains of Toxoplasma gondii. Vet. Parasitol., 68, 41, 1997.

Molecular Detection of Foodborne Pathogens 32. Jacobs, L., Remington, J.S., and Melton, M.L. A survey of meat samples from swine, cattle, and sheep for the presence of encysted Toxoplasma. J. Parasitol., 46, 23, 1960. 33. Dubey, J.P. Refinement of pepsin digestion method for isolation of Toxoplasma gondii from infected tissues. Vet. Parasitol., 74, 75, 1998. 34. Bell, A. and Ranford-Cartwright, L. Real-time quantitative PCR in parasitology. Trends Parasitol., 18, 338, 2002. 35. Contini, C. et al. Evaluation of a Real-time PCR-based assay using the lightcycler system for detection of Toxoplasma gondii bradyzoite genes in blood specimens from patients with toxoplasmic retinochoroiditis. Int. J. Parasitol., 35, 275, 2005. 36. Bastien, P. et al. Three years of multi-laboratory external quality control for the molecular detection of Toxoplasma gondii in amniotic fluid in France. Clin. Microbiol. Infect., 13, 430, 2007. 37. Calderaro, A. et al. Comparison between two real-time PCR assays and a nested-PCR for the detection of Toxoplasma gondii. Acta Biomed., 77, 75, 2006. 38. Burg, J.L. et al. Direct and sensitive detection of a pathogenic protozoan, Toxoplasma gondii, by polymerase chain reaction. J. Clin. Microbiol., 27, 1787, 1989. 39. Liesenfeld, O. et al. A case of disseminated toxoplasmosisvalue of PCR for the diagnosis. J. Infect., 29, 133, 1994. 40. Khalifa, K.-S. et al. Value of PCR for evaluating occurrence of parasitemia in immunocompromised patients with cerebral and extracerebral toxoplasmosis. J. Clin. Microbiol., 32, 2813, 1994. 41. Bretagne, S. et al. Quantitative competitive PCR with bronchoalveolar lavage fluid for diagnosis of toxoplasmosis in AIDS patients. J. Clin. Microbiol., 33, 1662, 1995. 42. Pelloux, H. et al. A second European collaborative study on polymerase chain reaction for Toxoplasma gondii, involving 15 teams. FEMS Microbiol. Lett., 165, 231, 1998. 43. Reischl, U. et al. Comparison of two DNA targets for the diagnosis of toxoplasmosis by real-time PCR using fluorescence resonance energy transfer hybridization probes. BMC Infect. Dis., 3, 7, 2003. 44. Contini, C. et al. The role of stage-specific oligonucleotide primers in providing effective laboratory support for the molecular diagnosis of reactivated Toxoplasma gondii encephalitis in patients with AIDS. J. Med. Microbiol., 51, 879, 2002. 45. Switaj, K. et al. Recent trends in molecular diagnostics for Toxoplasma gondii infections. Clin. Microbiol. Infect., 11, 170, 2005. 46. Goto, M. et al. Detection of Toxoplasma gondii by polymerase chain reaction in cerebrospinal fluid from human immunodeficiency virus-1-infected Japanese patients with focal neurological signs. J. Int. Med. Res., 32, 665, 2004. 47. Hurtado, A. et al. Single tube nested PCR for the detection of Toxoplasma gondii in fetal tissues from naturally aborted ewes. Vet. Parasitol, 102, 17, 2001. 48. Jauregui, L.H. et al. Development of a real-time PCR assay for detection of Toxoplasma gondii in pig and mouse tissues. J. Clin. Microbiol., 39, 2065, 2001. 49. Homan, W.L. et al. Identification of a 200- to 300-fold repetitive 529 bp DNA fragment in Toxoplasma gondii, and its use for diagnostic and quantitative PCR. Int. J. Parasitol., 30, 69, 2000. 50. Lin, M.H. et al. Real-time PCR for quantitative detection of Toxoplasma gondii. J. Clin. Microbiol., 38, 4121, 2000.

Toxoplasma 51. Kupferschmidt, O. et al. Quantitative detection of Toxoplasma gondii DNA in human body fluids by TaqMan polymerase chain reaction. Clin. Microbiol. Infect., 7, 120, 2001. 52. Darde, M.L., Bouteille, B., and Pestre-Alexandre, M. Isoenzyme analysis of 35 Toxoplasma gondii isolates and the biological and epidemiological implications. J. Parasitol., 78, 786, 1992. 53. Howe, D.K. and Sibley, L.D. Toxoplasma gondii comprises three clonal lineages: correlation of parasite genotype with human disease. J. Infect. Dis., 172, 1561, 1995. 54. Ajzenberg, D. et al. Genetic diversity, clonality and sexuality in Toxoplasma gondii. Int. J. Parasitol., 34, 1185, 2004. 55. Ajzenberg, D. et al. Microsatellite analysis of Toxoplasma gondii shows considerable polymorphism structured into two main clonal groups. Int. J. Parasitol., 32, 27, 2002. 56. Ajzenberg, D. et al. Genotype of 86 Toxoplasma gondii isolates associated with human congenital toxoplasmosis, and correlation with clinical findings. J. Infect. Dis., 186, 684, 2002. 57. Khan, A. et al. Recent transcontinental sweep of Toxoplasma gondii driven by a single monomorphic chromosome. Proc. Natl. Acad. Sci. USA, 104, 14872, 2007. 58. Lehmann, T. et al. Globalization and the population structure of Toxoplasma gondii. Proc. Natl. Acad. Sci. USA, 103, 11423, 2006. 59. Pena, H.F.J. et al. Population structure and mouse-virulence of Toxoplasma gondii in Brazil. Int. J. Parasitol., 38, 561, 2008. 60. Grigg, M.E. et al. Unusual abundance of atypical strains associated with human ocular toxoplasmosis. J. Infect. Dis., 184, 633, 2001. 61. Khan, A. et al. Genotyping of Toxoplasma gondii strains from immunocompromised patients reveals high prevalence of type I strains. J. Clin. Microbiol., 43, 5881, 2005. 62. Darde, M. Genetic analysis of the diversity in Toxoplasma gondii. Ann. Ist. Super Sanita., 40, 57, 2004. 63. Khan, A. et al. Composite genome map and recombination parameters derived from three archetypal lineages of Toxoplasma gondii. Nucleic Acids Res., 33, 2980, 2005. 64. Su, C., Zhang, X., and Dubey, J.P. Genotyping of Toxoplasma gondii by multilocus PCR-RFLP markers: A high resolution and simple method for identification of parasites. Int. J. Parasitol., 36, 841, 2006.

753 65. Dubey, J.P. et al. Biologic and genetic comparison of Toxoplasma gondii isolates in free-range chickens from the northern Para state and the southern state Rio Grande do Sul, Brazil revealed highly diverse and distinct parasite populations. V. Parasitol., 143, 182, 2007. 66. Ferreira, A.M. et al. Genetic analysis of natural recombinant Brazilian Toxoplasma gondii strains by multilocus PCRRFLP. Infect. Genet. Evol., 6, 22, 2006. 67. Ferreira, I.M. et al. Toxoplasma gondii: genotyping of strains from Brazilian AIDS patients with cerebral toxoplasmosis by multilocus PCR-RFLP markers. Exp. Parasitol., 118, 221, 2008. 68. Ajzenberg, D., Dumetre, A., and Darde, M.-L. Multiplex PCR for typing strains of Toxoplasma gondii. J. Clin. Microbiol., 43, 1940, 2005. 69. Blackston, C.R. et al. High-resolution typing of Toxoplasma gondii using microsatellite loci. J. Parasitol., 87, 1472, 2001. 70. Lehmann, T. et al. Variation in the structure of Toxoplasma gondii and the roles of selfing, drift, and epistatic selection in maintaining linkage disequilibria. Infect. Genet. Evol., 4, 107, 2004. 71. Sundar, N. et al. Genetic diversity among sea otter isolates of Toxoplasma gondii. Vet. Parasitol., 151, 125, 2008. 72. Sroka, J., Wójcik-Fatla, A., and Dutkiewicz, J. Occurrence of Toxoplasma gondii in water from wells located on farms. Ann. Agric. Environ. Med., 13, 169, 2006. 73. Costa, J.M. et al. Real-time PCR for diagnosis and follow-up of Toxoplasma reactivation after allogeneic stem cell transplantation using fluorescence resonance energy transfer hybridization probes. J. Clin. Microbiol., 38, 2929, 2000. 74. Howe, D.K. et al. Determination of genotypes of Toxoplasma gondii strains isolated from patients with toxoplasmosis. J. Clin. Microbiol., 35, 1411, 1997. 75. Nicoll, S. et al. A comparison of two methods of gene amplification for the diagnosis of Toxoplasma gondii in AIDS. J. Infect., 33, 177, 1996. 76. Prestrud et al. Direct high-resolution genotyping of Toxoplasma gondii in arctic foxes (Vulpes lagopus) in the remote arctic Svalbard archipelago reveals widespread clonal Type II lineage. Vet. Parasitol. 158,121, 2008.

Section VI Foodborne Helminthes

54 Anisakis

Stefano D’Amelio, Marina Busi, Sofia Ingrosso, and Lia Paggi Sapienza University of Rome

Elisabetta Giuffra

Parco Tecnologico Padano

Contents 54.1 Introduction.................................................................................................................................................................... 757 54.1.1 Classification . .................................................................................................................................................. 758 54.1.2 Biology and Epidemiology................................................................................................................................ 758 54.1.3 Pathogenesis and Medical Importance............................................................................................................. 759 54.1.4 Molecular Identification . ................................................................................................................................. 759 54.2 Methods...........................................................................................................................................................................761 54.2.1 Reagents and Equipment for Molecular Methods.............................................................................................761 54.2.2 Sample Preparation............................................................................................................................................761 54.2.2.1 Isolation of Larvae from Fish Hosts.................................................................................................761 54.2.2.2 DNA Purification..............................................................................................................................761 54.2.3 Detection Procedures........................................................................................................................................ 762 54.2.3.1 Primers............................................................................................................................................. 762 54.2.3.2 PCR Amplification ......................................................................................................................... 762 54.2.3.3 Restriction Fragment Length Polymorphisms Protocol.................................................................. 762 54.2.3.4 Identification Based on the ITS Restriction Patterns....................................................................... 762 54.2.3.5 PCR Product Purification and Sequencing...................................................................................... 762 54.2.3.6 Identification Based on the cox-2 Sequences................................................................................... 763 54.3 Conclusions and Future Perspectives............................................................................................................................. 763 References.................................................................................................................................................................................. 766

54.1  Introduction Anisakid nematodes are a homogeneous group constituted of species whose adult stages are recovered in fish, fish-eating birds, and marine mammals, while third stage larvae are present in the body cavity and muscle of numerous fish species. Anisakids are of medical and economic importance, due to public health implications and for the subsequent effects on the marketability of fish products. Larval forms of anisakid nematodes, in particular those belonging to the genera Anisakis and Pseudoterranova, are in fact the main causative agents of human anisakidosis, a fish-borne zoonosis due to the ingestion of raw or undercooked fish or cephalopods, that are infected by these larvae. Since the 1980s, pioneer studies on the genetic structure of anisakid nematodes have been carried out using allozyme markers, on a large number of taxa, sampled from many geographic regions from the Boreal and Austral hemispheres. Important achievements of these studies concern the detection of sibling species, the estimation of genetic divergence and of the relationships among taxa, the analysis of patterns of genetic variation in populations and species and measurement of

gene flow. In anisakid nematodes, cladogenetic events were accompanied by poor morphological differentiation, as ecological factors have led to a convergence to similar and well fitting morpho-functional solutions. This gave origin to a large number of morphologically identical species (sibling species). Therefore, morphological traits do not provide definitive evidence to be used as landmarks for their identification. This scarcity of diagnostic characters in adult individuals is even more remarkable in larval forms, where the number of structural traits usable for diagnostic purposes is more limited. Moreover it should be stressed that the larvae are responsible for human infection, and that their identification is essential for understanding the epidemiology of human anisakidosis. Also at higher taxonomic level, several morphological characters, even if strikingly differentiating one group from another, appear to be homoplastic and not always related to the phylogeny of species or genera. The poor taxonomic significance of some morphological characters and the occurrence of speciation processes without morphological differentiation undoubtedly advocate for the use of genetic and molecular approaches as efficient tools for the inference of the systematics and evolution of anisakid nematodes. 757

758

54.1.1 Classification The taxonomy of anisakids is mainly based on the morphology of male adult specimens. The most significant structural characters for species identification are the distribution and pattern of caudal papillae, the spicule and the morphology of cephalic end.1 Anisakid larvae can be identified at genus level by light microscopy, mainly on the basis of the morphology of digestive tract and of excretory system. By using multilocus allozyme electrophoresis, the analysis of the genetic structure of both adults and larvae of Anisakis species has provided interesting results at the taxonomic level.2–4 At present, allozyme, and PCR-RFLP studies have demonstrated that the genus Anisakis comprises at least nine distinct species, six belonging to Clade 1, corresponding to the larval Type I,5 namely, Anisakis simplex sensu stricto, Anisakis pegreffii, Anisakis simplex C, Anisakis typica, Anisakis ziphidarum, and Anisakis sp. A,6 which is identical to Anisakis sp. described by Valentini et al.7 and three belonging to Clade 2, showing the larval Type II,5 i.e., Anisakis physeteris, Anisakis brevispiculata, and Anisakis paggiae, which have different genetic structures, life cycles and geographical distributions.6,8–15 Recently, Palm et al.16 have hypothesized the existence of one additional taxa, closely related to A. typica, in a study carried out on Anisakis larvae collected in nonmigratory fish species in Balinese and Javanese waters, Indonesia. Finally, A. schupakovi, a parasite of the relict phocid species of the Caspian Sea, Phoca caspica, was considered as inquirendae by Davey17 in his revision of the genus Anisakis, but retained as a valid species by Kurochkin.18

54.1.2  Biology and Epidemiology Life cycle. Adult forms of the genera Anisakis and Pseudoterranova live in the stomach of marine mammals. Specifically members of the genus Anisakis are parasites of cetaceans, with the exception of A. schupakovi, parasite of a relict seal species in the Caspian Sea, Phoca caspica. As for the genus Pseudoterranova, the members of the P. decipiens complex are parasitic on seals. The life cycle begins when eggs are passed through the feces of the definitive host. Once the eggs are passed, they moult into second stage larvae. These must be consumed by an intermediate host, usually a crustacean. Predators of crustaceans, frequently fish or squid, become infected after eating an infected crustacean. Since anisakids do not undergo any development or moult inside the fish or squid, these hosts should be regarded as paratenic hosts of the nematode. The life cycle is completed after the paratenic host is ingested by a definitive host. It is possible that small fishes and squids are predated by larger fish species, thus determining an additional passage in a new paratenic host. Inside the stomach or the intestine of its final mammalian host, the worm develops into a sexually mature adult. While the above mentioned life cycle is accepted by many scientists, there is considerable evidence that two

Molecular Detection of Foodborne Pathogens

moults actually occur during development in the egg of A. simplex and therefore the third larval stage hatches from the egg. Host preference. Anisakid larvae have been detected worldwide in a large variety of fish species. Among teleosts, they have been found in Gadiformes, Perciformes, Clupeiformes, Pleuronectiformes, Scorpaeniformes, Zeiformes, Berici­ formes, Lophiiformes, Anguilliformes, and Atheriniformes. Anisakid larvae have been also detected in elasmobranchs and in a variety of cephalopods. Among these, they have been found in Octopodidae, Sepiidae, Loliginidae, and Ommastrephidae. In the stomach of cetaceans, anisakid adults are often found in clusters of individuals embedded in the mucosa and submucosa. Ulcers of 5 × 3 cm sized associated with anisakids are found in the fundic portion of the stomach. Six species have been grouped on the basis of the genetic relationships inferred from allozyme ad DNA sequence data, thus forming the Clade 1. A. simplex s.s. has been frequently found at its adult stage in Balaenoptera acutorostrata, Delphinapterus leucas, Delphinus delphis, Globicephala melaena, Lagenorhynchus albirostris, Orcinus orca, Pseudorca crassidens, Phocoena phocoena, and more rarely in Stenella coeruleoalba. Adults of A. pegreffii are frequent parasites of Delphinus delphis, Ziphius cavirostris and Tursiops truncates, and have been recorded also in Caperea marginata. The species A. simplex C has been identified as adult in the stomach of Lagenorhynchus albirostris, Lissodelphis borealis and Pseudorca crassidens. Two related species, Anisakis ziphidarum and Anisakis sp. A, are parasitic specifically of cetaceans belonging to the family Ziphiidae. In particular, adult forms of A. ziphidarum have been found in Ziphius cavirostris, Mesoplodon layardii, M. europaeus and M. densirostris. Anisakis sp. A (also described as Anisakis sp.) has been found as adult or preadult in Mesoplodon layardii, M. densirostris, M. mirus and M. grayi. The species A. typica has been recorded in several species of marine mammals. Mattiucci and Nascetti19,20 and D’Amelio et al.,21 have reported adult forms of this species in Globicephala macrorhynchus, Sotalia fluviatilis, Steno bredanensis, Stenella attenuata, S. longirostris, S. coeruleoalba, Tursiops truncatus, Lagenodelphis hosei, and Mesoplodon bidens. Three species of the genus Anisakis, strictly related one another, form the so called Clade 2. These species are A. physeteris, A. brevispiculata, and A. paggiae. The main definitive host for A. physeteris is the sperm whale, Physeter macrocephalus, but there are records of this species in the pigmy sperm whale, Kogia breviceps. A. brevispiculata, and A. paggiae have been recorded in Kogia breviceps and in the dwarf sperm whale, Kogia sima. They are often found in several fish species although showing a particular predilection for cephalopods, that represent important prey items of Physeter and Kogia species. Geographic distribution. Anisakis simplex s.s. has been recorded mainly in the North Atlantic and North Pacific Oceans, ranging from 35°N to the Arctic Circle. This species

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Anisakis

has been rarely found in the Mediterranean Sea, probably as a result of migration of pelagic fishes through the Gibraltar Strait. A. pegreffii is mainly distributed in the Mediterranean Sea and the South Atlantic Ocean. It is by far the most frequent species in the Mediterranean and the larvae of this species have been found in both pelagic and demersal fishes. Some records of this species have been reported in Japanese waters22 and New Zealand waters.19 The third species of the complex, A.simplex C, shows a “leopard skin” distribution, as it has been found in the Canadian and Californian Pacific waters, Chile, New Zealand and the Eastern South Atlantic Ocean.19 The species A. typica is generally found in warm temperate and tropical waters. Adults and larvae of this species have been found in several cetacean and fish species of the Gulf of Mexico, the Brazilian coasts of South America, coasts of Somalia, Mauritania, and Morocco and in the central-eastern part of the Mediterranean Sea, including Cyprus, Tunisia, and Lybia.23 The occurrence of A. typica in the eastern part of the Mediterranean has been related to migration of fishes from the Red Sea through the Suez Canal.19 A. ziphidarum has been found in the coasts of South Africa, the Mediterranean Sea, Central Atlantic waters up to the Carribean Sea, while its closely related species, Anisakis sp. A has been recorded in the Central Atlantic Ocean, in the coasts of Madeira and North-Western Spain, as well as in the waters of South Africa and New Zealand.6,24 A. physeteris is distributed in the Mediterranean Sea, including the coasts of North Africa,23 the Atlantic coasts of Spain, Central Atlantic Ocean, and the Gulf of Mexico. A. brevispiculata and A. paggiae show a geographic distribution along the warm-temperate waters of the Atlantic Ocean, from the Gulf of Mexico to the coasts of the Iberian Peninsula.9,10

54.1.3  Pathogenesis and Medical Importance Pathogenesis. Human anidakidosis, first reported in The Netherlands, has gained an increasing health and economic relevance in particular in countries such as Japan, where the consumption of raw fish and squids is frequent, although human cases are increasingly reported in the United States and many European countries (United Kingdom, France, Spain, and Italy). The anisakidosis of the digestive tract is classified from the clinical point of view, as acute or moderate. On the basis of the localization, the disease is subdivided in gastric anisakidosis and intestinal anisakidosis. The acute variety is prevalently gastric, and is characterized by nausea, vomiting, and epigastric pain. These symptoms appear 4–6 h after the ingestion of the infected fish. In the intestinal anisakidosis, acute signs appear after about 7 days after infection with abdominal pain, nausea, vomiting, fever, diarrhea, fecal occult blood, leukocytosis, and, rarely, eosinophilia. The

moderate forms of gastric anisakidosis are characterized by appetite loss, epigastralgia, and the occurrence of gastric pseudotumor; the evolution of moderate intestinal anisakidosis is more malicious. Several rare extragastrointestinal localizations have been documented (abdominal cavity, mesenteries, omentum). On the basis of the histological picture, the moderate variety of anisakidosis has been classified into four sequential stages: the first, or phlegmon, is typical of intestinal anisakidosis rather than gastric anisakidosis; the second, or abscess, is rather frequent gastric anisakidosis and is characterized by abundant necrotic tissue around the larvae and by a rich population of eosinophils; the third, or granulomaabscess, corresponds to the flogistic evolution of the disease, at least 6 months after the ingestion of the larva. At this stage the larva appears as few remnants invaded by the eosinophils, surrounded by giant cells and abundant inflammatory parvicellular infiltrate; the fourth, or granuloma stage is the most advanced, not frequently observed, characterized by a further decrease in the presence of eosinophils, with abundance of lymphocytes, giant cells, and significant collagenization. Allergies. Anisakidosis is also of medical importance because of the severe allergic reactions it causes in humans after eating or handling infected fish or crustaceans. These reactions include chronic urticaria (skin rashes), gastric ulcers, and anaphylaxis (a hyper-immune response). The allergic hypersensitivity symptoms in gastroallergic anisakidosis are clinical events accompanying a wide range of immunologic reactions as a host response against a ubiquitous parasite, but other frequent allergic disorders like chronic urticaria are now being studied for a possible relationship with A. simplex parasitism.25 Occupational hypersensitivity to A. simplex could occur in humans handling contaminated fish, e.g., in frozen-fish factories. Several case reports show allergy and anaphylactic reactions to the fish parasite Anisakis in the domestic and occupational settings. Diagnosis and treatment. The definitive diagnosis is often reached by endoscopic recovery of the larva and subsequent molecular identification at species level or by histological examination of endoscopic or surgical biopsies. The specificity of cutaneous tests (skin prick test) or serologic tests (ELISA, immunoblot) is still uncertain. The endoscopic removal of the larva in gastric anisakidosis seems to be resolutive of the disease. The effective use of antiheminthic drugs is not supported by large surveys, although recent studies have evidenced a significant action of tiabendazole and albendazole against these nematodes.

54.1.4 Molecular Identification The strong limit of the specific diagnosis of the etiological agents of anisakidosis on the basis of morphological examination relies mainly on two factors. The first is that larval forms of nematodes usually lack morphological

760

characters useful for taxonomic purposes. Anisakid nematodes do not represent an exception to this assumption. The larvae belonging to the genus Anisakis can be easily discriminated from the other genera of the family Anisakidae, and can be subdivided, on the basis of the presence/absence of the mucron, in two distinct larval types, but cannot be identified at species level, since each larval type is represented by more than one morphologically undistinguishable species. Moreover, the widespread phenomenon of the existence of sibling species in anisakids makes the identification at species level rather problematic even in adult male specimens, where the number of morphological characters usable for taxonomic purposes is definitively much higher. These factors make the use of genetic and molecular methods absolutely necessary to identify the larvae of anisakid nematodes at species level. The genetic, biochemical, and molecular methods so far applied in the taxonomy and molecular systematics of the genus Anisakis are summarized as follows: Allozymes. Multilocus allozyme electrophoresis has been extensively used in anisakid nematodes since the beginning of the 1980s to address questions concerning the population genetics, evolutionary biology, host-parasite co-evolution, and the relationship between genetic variability and habitat disturbance. The most important achievements of these studies are at the population and species levels, while above these taxonomic levels the methodology shows significant limitations. Among these results it is worth mentioning the detection of sibling species, i.e., species morphologically very similar, at times identical, but genetically and ecologically differentiated; the estimation of the amount of genetic divergence and of the relationships among closely related taxa; the analysis of patterns of genetic variation in populations and species and the measurement of gene flow within species.2–4,8,12,26–37 PCR-RFLP. Notwithstanding the huge bulk of data obtained so far from the application of multilocus allozyme electrophoresis, the development of molecular markers for the accurate identification of related species using PCR-based approaches may be in some cases preferable, considering that this approach has shown a remarkable sensitivity in the detection of genetic variation requiring only small amounts of fresh or ethanol-fixed parasite material for analysis. For example, PCR-based restriction fragment length polymorphism (PCR-RFLP) and sequence analyses of the ribosomal DNA (rDNA) internal transcribed spacers (ITS-1 and ITS-2) provide a useful approach for the specific identification of both distantly and closely related ascaridoid species, as these spacers show high levels of interspecific sequence differences in the presence of low-level intraspecific variation. PCR-based methods were therefore exploited to establish genetic markers in rDNA for the identification of anisakid nematodes belonging to the genera Anisakis (irrespective of developmental stages). In particular RFLP markers were obtained for the three sibling species of the A. simplex

Molecular Detection of Foodborne Pathogens

complex and for other four morphologically differentiated species of Anisakis, i.e., A. physeteris, parasite of sperm whales, A. typica, parasite of cetaceans in warm waters, A. ziphidarum, a parasite of cetaceans of the family Ziphiidae and A. schupakovi, parasite of the Caspian Seal, Phoca caspica.13 These markers have been successfully used in the identification of species of the genus Anisakis from fishes from Spain,14 from Madeira,6 from flatfish species from the coasts of Portugal,15 from the coasts of Japan,22 from the coasts of Tunisia,23 from migratory and non migratory fishes from Indonesia,16 from marine mammals from Galician coasts of Spain,24 and from the Atlantic coasts of Morocco and Mauritania.38 Multiplex and species-specific PCR. Umehara et al.39 have developed a method based on multiplex PCR able to selectively amplify six different species of anisakids including two species of the Anisakis simplex complex (A. simplex s.s. and A. pegreffii). The specificity of these primers to discriminate between the two sibling species was increased due to the introduction of artificial mismatched bases. Recently, PCR-sequence-specific primers were developed by Abe 40 in order to establish a quick method for the discrimination between A. simplex s.s. and A. pegreffii. DNA sequencing. Direct DNA sequencing proved a fruitful tool for the identification of the different sibling and morphospecies of Anisakis. Actually, sequence data are available for almost all the species recognized within the genus with the only exception of A. schupakovi. In particular the sequences available concern the region of the nuclear ribosomal DNA, spanning the final part of the 18S subunit, the first internal transcribed spacer (ITS-1), the 5.8S subunit, the second internal transcribed spacer (ITS-2) and the very beginning of the 28S subunit. The two ITS region are highly variable, in particular among different morphospecies, and show a significant degree of variation also among closely related species, as the three members of the A. simplex complex (i.e., A. simplex s.s., A. pegreffii and A. simplex C), useful for species discrimination. On the other hand, concerted evolution tends to minimize the intraspecific variation in this genomic region, thus allowing an unambiguous attribution of one specified sequence to one corresponding species. As for the mitochondrial DNA, two regions have been so far studied. The mitochondrial gene cox2 (cytochrome oxidase 2) was analyzed with the aim to clarify the phylogenetic relationships within the genus Anisakis.7 Although a certain degree of intraspecific variation could interfere with the use of this marker for diagnostic (not phylogenetic) purposes, in our opinion this region is highly reliable for species discrimination. The above markers have all been confirmed also by another mtDNA gene, the rrnS (the small subunit of the ribosomal DNA in the mitochondrial genome). Abollo et al.14 showed that the differences at this gene marker can be used to distinguish larvae of A. simplex s.s. and A. pegreffii.

761

Anisakis

54.2  Methods The entire protocol, from the isolation of larvae to their specific identification is here described, using two genetic markers based on the different nucleotide sequences in the species of the genus Anisakis. The most reliable markers are the internal transcribed spacers of the nuclear ribosomal DNA (ITS-1 and ITS-2) as developed by D’Amelio et al.13 and implemented by Pontes et al.6 and the mitochondrial gene coding for the cytochrome oxidase 2 (cox-2) as developed by Valentini et al.7 Based on nucleotide differences, a taxonomic key based on two diagnostic restriction enzymes (HhaI and HinfI) after amplification of the ITS genomic region has been developed for the identification of species of the genus Anisakis. Additionally, species identification can be achieved, or confirmed by direct sequencing of the PCR products after amplification using specific primers for the cox-2 mitochondrial gene.

54.2.1 Reagents and Equipment for Molecular Methods Reagents Wizard® SV Purification System NucleoSpin Extract II (MACHEREY-NAGEL) BIOTAQ™ DNA Polymerase Primers Restriction enzymes (Promega Hinf I 10 U/μl with Buffer B 10x and Hha I 10U/μl con Buffer C 10 × , BSA 10 mg/ml). Agarose TBE buffer Blue/orange loading dye Molecular marker (Hyperladder IV Bioline or 100bp DNA Ladder Promega)

Equipment Thermal bath Vortex Centrifuge Thermocycler Electrophoresis Gel Supply Chemical hood Gel Doc Microwave oven Micropipettes (20 μl, 100 μl, 200 μl, 1000 μl) Eppendorf tubes 1.5 ml PCR tubes 0.2 ml Flask 250 ml

54.2.2 Sample Preparation 54.2.2.1  Isolation of Larvae from Fish Hosts Anisakid larvae, due to their relatively large size, are easily recognizable from the body cavity or in the flesh and fillets of a large array of fish species and from some cephalopods. The larvae can be easily detected by ocular examination after opening the visceral cavity: After insertion of scissors in the anal opening of the fish, the operator proceed to a deep cut in tail-head direction up to the gills, plus two additional transversal cuts, as to allow a perfect view of the coelomatic cavity and of the mesentheric walls. It is convenient to remove the viscera in order to detect other larvae eventually present. The Anisakis larvae can be elongated or coiled, free or partially embedded on the surface of internal organs, such as liver and gonads.

Considering that the larvae tend to migrate in the flesh and fillets of fishes, especially after the death of the paratenic hosts, it appears needed an efficient examination of fish muscles. It is strongly suggested to use a transilluminator, although in larger fish, it could be necessary to isolate the larvae by muscular digestion using pepsin and chloridric acid. 54.2.2.2 DNA Purification We describe two different protocols to prepare DNA templates for PCR amplification. (i) In-house method: Procedure: (1) Introduce the larva in liquid nitrogen to be repeatedly frozen and thawed. (2) Pulverize the larva directly in a 1.5-ml plastic tube using a sterile pestle. (3) Add 100 µl of Holmes-Bonner solution (7 M Urea, 100 mM Tris-HCl pH 8.0, 10 mM EDTA pH 8.0, 350 mM NaCl, 2% SDS); and pestle to complete homogenization. (4) Add 100 µl of phenol-chloroform-isoamyl alcohol (50:49:1) and agitate for 10 min. (5) Centrifuge for 10 min at 13000 rpm. (6) Transfer the supernatant to a new sterile 1.5 ml tube. (7) Add 100 µl of chloroform and centrifuge for 5 min at 13000 rpm, then transfer the supernatant to a new sterile 1.5 ml tube. Repeat Step 7. (8) Add 200 µl of absolute ethanol, store at –20°C for 1h. (9) Centrifuge for 20 min at 13000 rpm, rinse with 200 µl of ethanol 70% and dry. (10) Resuspend pellet in 100 µl of TE containing RNase and store at room temperature for 30 min. (11) Run 5 µl on agarose gel to test. (ii) Wizard® SV Genomic DNA Purification System (Promega): Procedure: (1) Cut a 0.5–1.2-cm length of tissue or weigh up to 20mg of tissue sample and put it in a 1.5-ml microcentrifuge tube. (2) Add 275 μl of digestion solution master mix to each tube. Digestion Solution Master Mix Nuclei lisi solution

Volume Per Sample 200 μl

0.5 M EDTA (pH 8.0)

50 μl

Proteinase K 20 mg/ml

20 μl

RNase A solution 4 mg/ml Total volume

5 μl 275 μl

(3) Incubate the sample tubes overnight (16–18 h) in a 55°C heat block.

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(4) Add 250 μl of Wizard SV Lysis Buffer to each sample. Vortex. (5) Process the lysate as soon as possible after adding the Lysis Buffer. If frozen at –70°C, lysates should be thawed and heated at 55°C for 1 h prior to processing. Lysates must be warm for processing. Purification of genomic DNA from lysate using a microcentrifuge. (6) Transfer each sample lysate from the 1.5 ml tube to a separate Wizard SV Minicolumn Assembly. (7) Spin the assembly at 13000 × g for 3 min. (8) Remove minicolumn from the assembly and discard the liquid in the collection tube. (9) Replace the minicolumn into the collection tube. (10) Add 650 μl Wizard SV Wash Solution (with 95%ethanol added) to each assembly. Centrifuge at 13000 × g for 1 min. Discard the liquid from the collection tube. Repeat this step for a total of four washes. (11) Discard the liquid from the collection tube and reassemble the minicolumn assembly. Centrifuge for 2 min at 13000 × g to dry the banding matrix. (12) Transfer the Wizard SV Minicolumn to a new 1.5 ml tube. Add 250 μl of room temperature ­Nuclease-free water. Incubate for 2 min at room temperature. (14) Centrifuge the minicolumn/elution tube assembly at 13000 × g for 1 min. Do not discard the liquid in the elution tube. (15) Add an additional 250 μl of nuclease-free water and incubate at room temperature for 2 min. Centrifuge the minicolumn/elution tube assembly at 13000 × g for 2 min. (16) Remove the minicolumn and store the purified DNA at –20°C to –70°C.

54.2.3  Detection Procedures 54.2.3.1  Primers The entire ITS (ITS1, 5.8S rDNA gene and ITS2) can be amplified in species of the genus Anisakis using the primers NC5 (forward: 5′-GTAGGTGAACCTGCGGAAG GATCATT-3′) and NC2 (reverse: 5′- TTAGTTTCTTCCTCCGCT-3′). PCR amplification of the entire ITS in Anisakis species produces a fragment of about 1 kb. It is possible, for sequencing analysis, to separately amplify the two internal spacers (ITS-1 and ITS-2) using the following primers: The ITS-1 can be amplified in anisakid species using the primers NC5 (forward: 5′-GTAGGTGAACCTGCGGAAGGATCATT-3′) and NC13R (reverse: 5′-GCTGCGTTCTTCATCGAT-3′). PCR amplification of the ITS-1 in anisakid species produce a fragment of about 300 bp. The ITS-2 can be amplified in anisakid species using the primers XZ1 (forward: 5´-ATTGCGCCATCGGGTTCATTCC-3´) and NC2 (reverse: 5´-TTAGTTTCTTTTCCTCCGCT-3´). PCR amplification of the ITS-2 in anisakid species produces a fragment of about 400 bp. The cox-2 mitochondrial gene can be amplified using the

Molecular Detection of Foodborne Pathogens

primers 210 (forward: 5′-CACCAACTCTTAAAATTATC-3′) and 211 (reverse: 5′-TTTTCTAGTTATATAGATTGRTTY AT-3′). PCR amplification of the cox-2 in anisakid species produce a fragment of about 630 bp. Each primer must be diluted at 50 pmol/µl in sterile H2O. 54.2.3.2  PCR Amplification Prepare a PCR mix multiplying each reagent for the number of the individuals to be examined plus two sets for a positive and a negative control (e.g., for five specimens, prepare a complete set for the amplification of seven samples) as follows: (1) Add sequentially, for each sample, 5 µl 10 × PCR buffer, 5 µl MgCl2, 4 µl dNTPs, 0.5 µl of each primer, 0.3 µl Taq DNA polymerase and sterile H2O up to 48 µl in a sterile 1.5 ml tube. (2) Add 2 µl of genomic DNA to each 0.2 ml thinwalled tube. (3) Add 48 µl of PCR mix to each 0.2 ml thin-walled tube. Place tubes on ice. (4) PCR cycle: one cycle of 95°C for 10 min, 30 cycles of 95°C for 30 sec, 55°C for 30 sec and 72°C for 75 sec, and one cycle of 72°C for 7 min. Place tubes on ice. (5) Electrophoresis: load 5 µl of each amplification reaction to check the quality of amplification products and the fragment size on a 1% agarose gel containing 0.5 µg/ml ethidium bromide in TBE buffer at 10 V/cm. 54.2.3.3 Restriction Fragment Length Polymorphisms Protocol (1) Add sequentially, up to a final volume of 15.2 µl, 10 µl of PCR-amplified DNA, 3 µl of distilled water, 0.5 µl of restriction enzyme (HinfI and HhaI), 1.5 µl of enzyme buffer and 0.2 µl of BSA. (2) Incubate for 90 min at 37°C (with the exception of the endonuclease TaqI, to be incubated at 65°C). (3) Electrophoresis: load all the reaction on a 2% agarose gel containing 0.5 µg/ml ethidium bromide in TBE buffer at 10 V/cm. 54.2.3.4 Identification Based on the ITS Restriction Patterns Following the visualization on the gene of the PCR products after digestion with two restriction enzyme (HinfI, and HhaI) a useful DNA marker, the identification of larvae or adult at species level is made by comparison with the expected restriction patterns reported in Table 54.1. 54.2.3.5  PCR Product Purification and Sequencing PCR product purification. The cox-2 amplified fragments will be purified and subjected to direct sequencing. The purification protocol is as follows: The “NucleoSpin Extract II

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Anisakis

Table 54.1 Expected Restriction Patterns After Digestion of the ITS Amplified Fragments Using the HinfI and HhaI Endonucleases Fragment Sizes (bp) HinfI

HhaI

A.simplex s.s. A. pegreffii A. simplex C A. typica

620-250-80 370-300-250 620-250-80 610-350

550-430 550-430 550-300-130 320–240–180–160

A. ziphidarum Anisakis sp. A A. physeteris A. brevispiculata A. paggiae

370-320-290 620-320 380–290–270 1000 1000

550-430 550-300-80 540–420 400-320-200  520-400

Species

(MACHEREY-NAGEL) is suitable for this purpose. The purification protocol is as follows: (1) Adjust DNA binding conditions. Mix 2 volumes of buffer NT with 1 volume of sample (e.g., 200 μl and 100 μl PCR reaction mix). For sample volumes < 50 μl adjust the volume of the reaction mix to 50 μl using TE buffer pH 7.5. (2) Bind DNA. Place a NucleoSpin® Extract II column into a 2 ml collecting tube and load the sample. Centrifuge for 1 min at 11000 × g. Discard flowthrough and place the NucleoSpin® Extract II column back into the collecting tube. (3) Wash silica membrane. Add 600 μl buffer NT3. Centrifuge for 1 min at 11000 × g. Discard flowthrough and place the NucleoSpin® Extract II column back into the collecting tube. (4) Dry silica membrane. Centrifuge for 2 min at 11000 × g to remove buffer NT3 quantitatively. Note. Make sure the spin column doess not come in contact with the flow-through while removing it from the centrifuge and the collecting tube. Residual ethanol from buffer NT3 would inhibit subsequent reactions and has to be removed in this step. In addition to centrifugation, total removal can be achieved by incubation of NucleoSpin® Extract II columns for 2–5 min at 70°C prior to elution. (5) Elute DNA. Place the NucleoSpin® Extract II column into a clean 1.5 ml microcentrifuge tube. Add 15–50 μl elution buffer NE, incubate at room temperature for 1 min to increase the yield of eluted DNA. Centrifuge for 1 min at 11000 × g. DNA sequencing. Several services for DNA sequencing are available. It can be, in some conditions, more cost effective to use these services rather than perform sequencing in your own lab. Therefore a protocol for DNA sequencing is not included. Most sequencing services provide sequence results

in different modalities including text files, pdf files, and electropherograms. 54.2.3.6  Identification Based on the cox-2 Sequences The DNA sequences of the cox-2 mitochondrial gene can be used for species identification by direct comparison with the available sequences deposited in GenBank. The alignment of these sequences (including their Accession Numbers) is reported in Figure 54.1. This method is built upon the basis of the paper published by Valentini et al.7

54.3 Conclusions and Future Perspectives The systematics and classification of ascaridoid nematodes and the relationships among genera and species belonging to this superfamily have been the subject of many speculations. The proposed classifications, based mostly on the analysis of morphological traits considered of high systematic significance, are far from unambiguous and are to some extent inconsistent. Since the 1980s studies on the genetic structure of anisakid nematodes have been carried out on a large number of taxa, sampled from many geographic regions from the Boreal and Austral hemispheres. Important achievements of these studies concern the detection of sibling species, the estimation of genetic divergence and of the relationships among taxa, the analysis of patterns of genetic variation in populations and species and measurement of gene flow. In anisakid nematodes, cladogenetic events were accompanied by poor morphological differentiation, as ecological factors have led to a convergence of similar and well fitting morphofunctional solutions. This gave origin to a large number of morphologically identical species (sibling species). Also at higher taxonomic level, several morphological characters, even if strikingly differentiating one group from another, appear to be homoplastic and not related to the phylogeny of species or genera. Therefore, morphological traits do not provide definitive evidence to be used as landmarks for the inference of anisakid evolutionary histories. The poor phylogenetic significance of some morphological characters and the occurrence of speciation processes without morphological differentiation undoubtedly advocate for the use of genetic and molecular approaches as efficient tools for the inference of the systematics and evolution of anisakid nematodes.41 The discovery of new species in anisakid nematodes seems to be a continuous process, as the number of new sibling species detected by molecular methods increases as new geographical regions or new hosts are studied. This makes the systematics of the genus Anisakis and of related anisakid genera in a state of frequent rearrangement. The main objective of the development of molecular methods for the characterization of anisakid nematodes, and in particular of the species of the genus Anisakis, has been so far to provide a quick and reliable tool for the identification of specimens at species level, irrespective of their sex or life history stage. This has had a strong impact on the systematics

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Molecular Detection of Foodborne Pathogens

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

TTT TTT TTT TTT TTT TTT TTT TTT TTT

TCT TCT TCT TCT TCT TCT TCT TCT TCT

AGT AGT AGT AGT AGT AGT AGT AGT AGT

TAT TAT TAT TAT TAT TAT TAT TAT TAT

ATA ATA ATA ATA ATA ATA ATA ATA ATA

GAT GAT GAT GAT GAT GAT GAT GAT GAT

TGG TGG TGG TGG TGG TGG TGG TGG TGG

TTT TTT TTT TTT TTT TTT TTT TTT TTT

CAT CAT CAT CAT CAT CAT CAT CAT CAT

AAC AAC AAC AAT AAT AAT AAT AAG AGT

TTT TTT TTT TTT TTT TTT TTT CTT TTT

AAT AAT AAT AAT AAT AAT AAT TAT AAT

TGT TGT TGT TGT TGT TGT TGT TGT TGT

AGT AGT AGT AGT AGT AGT AGG AGG AGG

TTG CTG TTG TTG TTG TTG TTG TTA TTG

CTT CTT CTT CTT CTT CTT CTA CTT CTT

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

TTT TTT TTT TTT TTT TTT TTT TTT TTT

GGT GGT GGT GGT GGT GGT GGT GGT GGG

GTT GTT GTT GTT GTG GTC GTA GTG GTG

CTT CTT CTT TTA TTA TTG TTG TTA TTG

TCT TCT TCT TCT TCA TCT GCT GCT GCT

TTT TTT TTT TTT TTT TTT TTT TTT TTT

GTT GTA GTT GTT GTT GTC GTT GTT GTT

TCT TCT TCT TCT TCT TCT ACG ACA TCT

GTT GTT GTT GTT GTT GTT GTT GTT GTT

ATG ATG ATG ATA ATG ATG ATA ATA ATA

TTT TTT TTT TTT TTT TTT TTT TTT TTT

GGT GGC GGC GTT GTT GTA GTT GTT GTT

TAT TAT TAT TAT TAT TAC TAT TAT TAT

CTT CTT CTT TTA TTA TTG TTA TTA TTA

CTT CTT CTT CTT CTT CTT CTT CTT CTT

TTT TTT TTT TTT TTT TTT TTT TTT TTT

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

AGA AGA AGA AGG AGC AGC AAT AAT AAT

AAT AAT AAT AGT AGT AGT AGT AGG AGT

TTT TTT TTC TTT TTT TTT TTT TTT TTT

TAT TAT TAT TAT TAT TAT TAT TAT TAT

TTT TTT TTT TTT TTT TTT TTT TTT TTT

AAG AAA AAA AAG AAG AAG AAG AAA AAA

AGT AGT AGT AGA AGG AGG AGT AGT AGT

AAG AAG AAG AAG AAA AAG AAA AAA AAA

AAG AAG AAG AAG AAG AAG AAA AAG AAG

ATT ATT ATT ATT ATT ATT ATT ATT ATT

GAA GAG GAA GAA GAG GAG GAG GAG GAA

TAT TAT TAT TAT TAT TAT TAT TAT TAT

CAG CAG CAG CAG CAG CAG CAG CAG CAG

TTT TTT TTT TTT TTT TTC TTT TTT TTT

GGT GGT GGT GGT GGT GGG GGT GGT GGT

GAA GAA GAG GAA GAA GAA GAA GAG GAA

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

CTT CTT CTT CTT CTT CTT CTT CTT CTT

TTA TTA TTA TTG TTG TTG TTA CTT TTG

TGT TGT TGT TGT TGC TGT TGT TGT TGC

AGT AGT AGT AGT AGT AGT AGT AGT AGT

ATT ATT ATT ATT ATT ATT GTG GTC ATT

TTT TTT TTT TTC TTT TTT TTT TTT TTT

CCT CCT CCT CCT CCT CCT CCT CCT CCT

ACT ACT ACT ACT ACT ACT ACT ACT ACT

TTA TTA TTA CTT TTA TTA TTG TTG TTA

ATT ATT ATT ATT ATT ATT ATT ATC ATT

TTG TTG TTG TTG TTG CTG TTG TTA TTG

GTT GTT GTT GTG GTG GTG GTA GTT GTG

GCT GCT GCT GCC GCC GCC GCT GCT TTT

CAG CAG CAG CAA CAA CAG CAG CAA CAG

ATG ATG ATG ATG ATG ATA ATG ATA ATG

GTG GTG GTG GTG GTG GTG GTG GTA GTT

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

CCT CCT CCT CCT CCC CCC CCT CCT CCT

TCT TCT TCT TCT TCT TCT TCT TCT TCT

TTG TTG TTG TTA TTA TTG TTA TTA TTA

AGT AGT AGT AGT AGT AGA AGA AGA AGT

TTA TTA CTA TTG TTG TTG CTT CTT CTT

CTT CTT CTT CTT CTT CTT TTG CTT CTT

TAT TAT TAT TAT TAT TAT TAT TAC TAC

TAT TAT TAC TAT TAT TAT TAT TAT TAT

TAT TAT TAT TAT TAT TAT TAT TAT TAT

GGT GGT GGT GGT GGT GGT GGT GGT GGT

TTG TTG CTA TTG TTG TTA CTT TTG TTA

ATG ATG ATG ATA ATA ATA ATG ATG ATA

AAT AAT AAT AAT AAT AAT AAT AAT AAT

CTT CTT CTT TTG TTG CTT CTT CTT TTG

GAT GAT GAT GAT GAT GAT GAT GAT GAT

AGT AGT AGT AGT AGT AGT AGT AGG AGA

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris

AAT AAT AAT AAT AAT AAT AAT

TTG TTA TTA TTG TTA TTA CTT

ACT ACT ACT ACT ACG ACT ACT

GTG GTT GTA GTT GTA GTC GTT

AAA AAA AAA AAA AAG AAG AAG

GTT GTT GTT GTT GTA GTT GTT

ACT ACT ACG ACT ACT ACA ACT

GGT GGT GGC GGT GGG GGC GGT

CAT CAT CAT CAT CAT CAC CAT

CAG CAG CAG CAA CAG CAG CAA

TGG TGG TGG TGA TGA TGG TGG

TAT TAT TAC TAT TAT TAT TAT

TGG TGG TGG TGG TGG TGG TGA

AGT AGT AGT AGT AGT AGA AGA

TAT TAT TAT TAT TAT TAT TAT

GAG GAG GAG GAG GAA GAG GAA

Figure 54.1  Alignment of nucleotide sequences available for nine species of the genus Anisakis for the cox-2 mitochondrial gene. The corresponding accession numbers are as follows: A. simplex s.s.: DQ116426, A. pegreffii: DQ116428, A. simplex C: DQ116429, A. ziphidarum: DQ116430, Anisakis sp. A: DQ116431, A. typica: DQ116427, A. physeteris: DQ116432, A. brevispiculata: DQ116433, A. paggiae: DQ116434. (From Valentini, A. et al., J. Parasitol., 92, 156, 2006.)

765

Anisakis

A.brevispiculata A.paggiae

AAC TTG ACT GTT AAA GTA ACT GGT CAT CAA TGA TAT TGA AGG TAT GAG AAC TTA ACT GTT AAA GTT ACT GGT CAT CAA TGA TAT TGA AGT TAT GAG

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

TTT TTT TTT TTT TTT TTT TTT TTT TTT

AGT AGG AGG AGT AGT AGT AGA AGT AGT

GAT GAT GAT GAT GAT GAT GAT GAT GAT

ATC ATC ATT ATT ATT ATC ATT ATT ATT

CCG CCG CCT CCT CCC CCT CCT CCT CCC

GGY GGC GGT GGT GGA GGG GGT GGG GGT

TTA TTA TTG TTG TTG TTA TTA CTA TTA

GAA GAA GAA GAG GAG GAG GAA GAG GAA

TTT TTT TTT TTT TTT TTT TTT TTT TTT

GAT GAT GAT GAT GAT GAT GAT GAT GAC

TCT TCT TCT TCA TCT TCT TCT TCT TCT

TAT TAT TAT TAT TAT TAT TAT TAT TAT

ATG ATG ATG ATG ATA ATG ATG ATA ATG

AAG AAG AAG AAG AAG AAG AAG AAA AAG

TCT TCT TCT TCT TCT TCT TCT TCG TCT

GTG GTG GTA GTG GTT GTT GTG GTG GTA

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

GAT GAC GAT GAT GAT GAT GAT GAT GAT

CAG CAG CAG CAG CAA CAG CAG CAG CAG

TTG TTG TTA TTG TTG TTA TTA CTG TTG

GAG GAG GAG GAG GAG GAG GAG GAG GAG

CTA CTA TTA TTG TTG TTG TTG TTG TTA

GGT GGT GGT GGA GGC GGT GGA GGT GGA

GAG GAG GAG GAG GAG GAG GAG GAG GAG

CCT CCT CCT CCC CCC CCT CCA CCA CCA

CGT CGT CGT CGT CGT CGT CGT CGT CGT

CTT CTT CTT CTG TTG CTG CTT CTT CTA

TTG TTG TTG CTT TTG TTG TTA TTA CTA

GAG GAG GAG GAG GAA GAG GAG GAG GAG

GTT GTT GTT GTG GTT GTT GTT GTG GTT

GAT GAT GAT GAT GAT GAT GAT GAT GAT

AAT AAT AAT AAT AAC AAC AAT AAT AAT

CGT CGT CGT CGT CGT CGT CGT CGT CGT

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

TGT TGT TGT TGT TGT TGT TGT TGT TGT

GTT GTC GTT GTA GTT GTT GTG GTG GTG

GTT GTT GTT GTT GTT GTG GTT GTT GTT

CCT CCT CCT CCT CCT CCT CCT CCT CCG

TGT TGT TGT TGT TGT TGT TGT TGT TGT

GAT GAT GAT GAT GAT GAT GAT GAT GAT

ATT ATT ATT ATT ATT ACT ATT ATT ATT

AAT AAT AAT AAT AAT AAT AAC AAT AAT

GTC GTT GTG ATC ATT ATT ATT ATT ATT

CGT CGT CGT CGT CGT CGT CGT CGT CGT

TTT TTT TTT TTT TTT TTC TTT TTT TTC

TGT TGT TGT TGT TGT TGT TGT TGT TGT

ATT ATT ATT ATT ATT ATC ATT ATT ATT

ACC ACT ACT ACT ACT ACT ACT ACT ACT

TCG TCG TCT TCT TCC TCG TCG TCC TCT

GGG GGG GGG GGG GGG GGG GGT GGG GGC

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

GAT GAT GAC GAT GAT GAT GAT GAT GAT

GTT GTT GTT GTT GTT GTT GTA GTA GTT

ATT ATT ATT ATC ATT ATT ATT ATT ATT

CAT CAT CAT CAT CAT CAC CAT CAT CAT

TCT TCT TCT TCT TCT TCT TCT TCT TCT

TGG TGG TGG TGG TGG TGG TGG TGG TGG

GCT GCT GCT GCT GCT GCT GCT GCT GCT

TTG TTG TTG TTG TTG TTG TTG TTA TTG

CCT CCT CCG CCT CCC CCT CCT CCC CCT

AGA AGG AGA AGG AGA AGT AGA AGG AGT

ATA ATG ATG ATG ATG ATG ATG ATG ATG

TCT TCT TCT TCT TCT TCT TCT TCT TCT

ATT ATT ATT ATT ATT ATT ATT ATT ATT

AAG AAG AAG AAA AAG AAG AAA AAA AAA

TTG TTG TTG TTG TTG CTG CTT CTT CTT

GAT GAT GAT GAT GAT GAT GAT GAT GAT

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

GCT GCC GCT GCT GCT GCT GCT GCT GCT

ATA ATG ATA ATG ATG ATG ATG ATA ATA

AGG AGT AGG AGA AGA AGG AGG AGA AGA

GGT GGT GGT GGT GGT GGG GGT GGT GGT

ATT ATT ATT ATT ATC ATC ATT ATT ATT

TTG TTA TTG TTG TTA CTT TTG TTG TTA

TCT TCT TCT TCT TCT TCT TCT TCT TCT

ACT ACT ACT ACT ACA ACT ACT ACT ACT

GTT GTT GTT TTG TTA CTA TTA TTG TTG

TCT TCT TCT TCT TCT TCT TCT TCT TCT

TAT TAT TAT TAT TAC TAT TAT TAT TAT

AGT AGT AGT AGT AGT AGT AGT AGC AGT

TTT TTT TTT TTT TTT TTT TTT TTT TTT

CCT CCT CCT CCT CCT CCT CCA CCA CCT

ACT ACT ACT ACT ACT ACT ACT ACT ACT

GTT GTA GTT GTG GTG GTG GTG GTG GTA

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris

GGT GGT GGT GGG GGT GGA GGG

GTG GTG GTT GTT GTT GTT GTT

TTT TTT TTT TTT TTT TTT TTT

TAT TAT TAT TAT TAT TAC TAT

GGT GGT GGT GGT GGT GGG GGT

CAA CAA CAA CAG CAG CAA CAA

TGT TGT TGT TGC TGC TGT TGT

TCA TCA TCA TCG TCA TCT TCT

GAG GAG GAA GAG GAG GAA GAG

ATT ATT ATT ATC ATT ATT ATT

TGT TGT TGT TGT TGT TGT TGT

GGG GGG GGG GGG GGT GGG GGG

GCT GCT GCT GCT GCT GCT GCT

AAC AAC AAT AAT AAT AAT AAT

CAT CAT CAT CAT CAT CAT CAT

AGT AGT AGA AGT AGT AGT AGT

Figure 54.1  (Continued)

766

Molecular Detection of Foodborne Pathogens

A.brevispiculata A.paggiae

GGG GTT TTT TAT GGT CAA TGT TCT GAA ATT TGT GGG GCT AAT CAT AGT GGT GTT TTT TAT GGT CAA TGT TCT GAA ATT TGT GGA GCT AAC CAT AGT

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

TTT TTT TTT TTT TTT TTT TTT TTT TTT

ATG ATG ATG ATG ATG ATG ATA ATA ATG

A.simplex s.s. A.pegreffii A.simplex C A.ziphidarum Anisakis sp. A A.typica A.physeteris A.brevispiculata A.paggiae

TGG TGG TGG TGG TGG TGG TGG TGG TGG

TG TG TG TG TG TG TG TG TG

CCT CCC CCT CCC CCT CCT CCT CCT CCT

ATT ATT ATT ATT ATT ATT ATT ATT ATT

GCT GCT GCT GCT GCT GCT GCT GCT GCT

TTG TTG TTG TTA TTG TTA TTG TTG TTA

GAA GAA GAA GAG GAG GAG GAA GAG GAG

GTG GTG GTG GTG GTT GTA GTA GTA GTA

ACT ACT ACT ACT ACT ACT ACT ACT ACT

TTA TTG TTG TTG TTG TTG CTT TTA CTA

TTG TTG CTG CTT CTT CTT CTT CTT CTT

GAT GAT GAT GAT GAT GAT GAT GAT GAT

AAT AAT AAT AAT AAT AAT AAT AAT AAT

TTT TTT TTT TTT TTT TTT TTT TTT TTT

AAG AAG AAG AAG AAG AAG AAG AAG AAG

AGT AGT AGT AGT AGT AGT AGT AGT AGT

Figure 54.1  (Continued)

of the genus and on the ecology of the different species and has provided to physicians an effective method for the identification of the larvae recovered after endoscopy. Only a few cases in which the nematodes obtained from human infections have been identified at species level, using nuclear42–44 or mitochondrial,45 markers have been so far reported. The identification of the etiological agent can aid physicians in diagnosis and treatment, in tracing the source of the infestation, in determining the area of origin of the contaminated food, and in developing the most appropriate measures for controlling contamination at all phases of food production, from fishing, or breeding to processing and post­processing of fish products. However, these investigations on the identification of the larvae from fish and squids are based on recovered larvae after examination by eye and do not represent a significant step forward in terms of the detection of anisakid larvae in the fish hosts or in secondary fish products, both as entire larva or as a portion of it. On the other hand the occurrence of a small portion of parasite material in fishes, even if adequate freezing and/or cooking can kill or inactivate anisakid larvae, can still cause sensitization and IgE-dependent hypersensitivity in humans. Also non-IgE mediated mechanisms, such as the involvement of other immunoglobulin isotypes (IgG4), or nonimmunological events have been described.46 Heat- and/or pepsin-­resistant allergens from A. simplex could explain reactions and symptoms after the ingestion of well-cooked or canned fish.47 In this context it appears necessary to devote efforts in the direction of establishing direct methods for the detection of the parasite and/or of parasite material in a variety of products, either fresh or frozen, e.g., canned fish, “surimi” (minced, processed fish used in the preparation of imitation seafood), fish

sticks, and baby food (easily chewed food products for infants produced in multiple varieties and tastes, and produced by many manufacturers). For these derived products, no methods are currently available to detect and quantify the presence of larvae residuals that still carry allergen properties. Recently, a project has been designed in order to provide a new method for the detection and quantification of nematode parasites (Anisakis spp. and Pseudoterranova spp.) typically contaminating fishery species, and consequently fish products, by means of a real-time PCR assay which exploits the conserved features of the ITS-1 region of the Anisakid genome, optimized on standardized DNA samples. The preliminary results of this project, still unpublished and currently under patenting, have demonstrated that realtime PCR assay is a reliable method for the detection of residual anisakid DNA, not only in fish but also in secondary fish products. Considering that the existing regulations at EU level necessitates freezing fish or fish products to be consumed uncooked, as a preventative measure, useful to avoid the classical form of the disease, will be absolutely ineffective with regard to the allergenic properties of small residuals of the larvae. This makes the real-time PCR method one of the most sensitive and promising approaches for a “true” molecular detection of the parasite and also for its quantification. Moreover the probes used in the real-time PCR assay can be designed in species-specific portion of the ITS sequence, thus allowing the detection and the identification in the same assay.

References

1. Fagerholm, H.P. Systematic implications of male caudal ­morphology in ascaridoid nematode parasites. Syst. Parasitol., 19, 215, 1991.

Anisakis























2. Mattiucci, S. et al. Genetic structure of Anisakis physeteris, and its differentiation from the Anisakis simplex complex (Ascaridida: Anisakidae). Parasitology, 93, 83, 1986. 3. Mattiucci, S. et al. Genetic and ecological data on the Anisakis simplex complex, with evidence for a new species (Nematoda, Ascaridoidea, Anisakidae). J. Parasitol., 86, 401, 1997. 4. Nascetti, G. et al. Electrophoretic studies on the Anisakis simplex complex (Ascaridida: Anisakidae) from the Mediterranean and North-East Atlantic. Int. J. Parasitol., 16, 633, 1986. 5. Berland, B. Nematodes from some Norwegian marine fishes. Sarsia 2, 1, 1961. 6. Pontes, T. et al. Molecular characterization of larval anisakid nematodes from marine fishes of Madeira by a PCR-based approach, with evidence for a new species. J. Parasitol., 91, 1430, 2005. 7. Valentini, A. et al. Genetic relationships among Anisakis species (Nematoda: Anisakidae) inferred from mitochondrial cox2 sequences, and comparison with allozyme data. J. Parasitol., 92, 156, 2006. 8. Mattiucci, S. et al. Allozyme and morphological identification of Anisakis, Contracaecum and Pseudoterranova from Japanese waters (Nematoda, Ascaridoidea). Syst. Parasitol., 40, 81, 1998. 9. Mattiucci, S. et al. Genetic divergence and reproductive isolation between Anisakis brevispiculata and Anisakis physeteris (Nematoda: Anisakidae). Int. J. Parasitol., 31, 9, 2001. 10. Mattiucci, S. et al. Genetic markers in the study of Anisakis typica (Diesing, 1860): larval identification and genetic relationships with other species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae). Syst. Parasitol., 51, 159, 2002. 11. Mattiucci, S. et al. Evidence for a new species of Anisakis Dujardin, 1845: morphological description and genetic relationships between congeners (Nematoda: Anisakidae). Syst. Parasitol., 61, 157, 2005. 12. Paggi, L. et al. A new species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae) from beaked whales (Ziphiidae): allozyme and morphological evidence. Syst. Parasitol., 40, 161, 1998. 13. D’Amelio, S. et al. Genetic markers in ribosomal DNA for the identification of members of the genus Anisakis (Nematoda: Ascaridoidea) defined by polymerase chain reaction based restriction fragment length polymorphism. Int. J. Parasitol., 30, 223, 2000. 14. Abollo, E. et al. Occurrence of recombinant genotypes of Anisakis simplex s.s. and Anisakis pegreffii (Nematoda: Anisakidae) in an area of sympatry. Infect. Genet. Evol., 3, 175, 2003. 15. Marques, J.F. et al. Molecular identification of Anisakis species from Pleuronectiformes off the Portuguese coast. J. Helminthol., 80, 47, 2006. 16. Palm, H. W. et al. Molecular genotyping of Anisakis Dujardin, 1845 (Nematoda: Ascaridoidea: Anisakidae) larvae from marine fish of Balinese and Javanese waters, Indonesia. Helminthologia, 45, 3, 2008 17. Davey, J.T. A revision of the genus Anisakis Dujardin, 1845 (Nematoda: Ascaridida). J. Helminthol., 45, 51, 1971. 18. Kurochkin, Y.V. Parasites of the Caspian seal Pusa caspica. Rapport et proces-verbaux des reunion. Cons. Int. Expl. Mer., 169, 363, 1975. 19. Mattiucci, S. and Nascetti, G. Molecular systematics, phylogeny and ecology of anisakid nematodes of the genus Anisakis Dujardin, 1845: an update. Parasite, 13, 99, 2006.

767 20. Mattiucci, S. and Nascetti, G. Genetic diversity and infection levels of anisakid nematodes parasitic in fish and marine mammals from Boreal and Austral hemispheres. Vet. Parasitol., 148, 43, 2007. 21. D’Amelio, S. et al. Molecular genotyping of anisakid nematodes from cetaceans of Florida waters. European Cetacean Society Annual Conference, Cetacean Social Organization. La Rochelle (France) 2–7 April, 2005. 22. Umehara, A. et al. Molecular identification of Anisakis simplex sensu stricto and Anisakis pegreffii (Nematoda: Anisakidae) from fish and cetacean in Japanese waters. Parasitol. Int., 55, 267, 2006. 23. Farjallah, S. et al. Occurrence and molecular identification of Anisakis spp. from the North African coasts of Mediterranean Sea. Parasitol. Res., 102, 371, 2008. 24. Iglesias, R. et al. Molecular and morphological evidence of a new taxon of Anisakis Dujardin, 1845 (Nematoda, Anisakidae) from the Blainville’s beaked whale (Mesoplodon densirostris). J. Helminthol., 82, 305, 2008. 25. Daschner A. and Pascual C.Y. Anisakis simplex: sensitization and clinical allergy. Curr. Opin. Allergy Clin. Immunol., 5, 281, 2005. 26. Paggi, L. et al. Biochemical taxonomy of ascaridoid nematodes. Parassitologia, 27, 105, 1985. 27. Paggi, L. et al. Genetic evidence for three species within Pseudoterranova decipiens (Nematoda, Ascaridida, Ascari­ doidea) in the North Atlantic and Norwegian and Barents Seas. Int. J. Parasitol., 21, 195, 1991. 28. Paggi, L. et al. Nematodi del genere Anisakis in pesci, cefalopodi e cetacei del Mar Mediterraneao e dell’Oceano Atlantico e Pacifico. Biol. Mar. Medit., 5, 1585, 1998. 29. Paggi, L. et al. Pseudoterranova decipiens species A and B (Nematoda, Ascaridoidea): nomenclatural designation, morphological diagnostic characters and genetic markers. Syst. Parasitol., 45, 185, 2000. 30. Bullini, L., et al. Genetic variation of ascaridoid worms with different life cycles. Evolution, 40, 437, 1986. 31. Bullini, L. et al. Genetic and ecological studies on nematode endoparasites of the genera Contracaecum and Pseudoterranova in the Ant­arctic and Arctic-Boreal regions. Proceedings of the 2nd Meeting on “Biology in Antarctica”. Scienza e Cultura. Numero Speciale. Edizioni Universitarie Patavine, 131, 1994. 32. Nascetti, G. et al., Three sibling species within Contracaecum osculatum (Nematoda, Ascaridida, Ascaridoidea) from the Atlantic Arctic-Boreal region: reproductive isolation and host preferences, Int. J. Parasitol., 23, 105, 1993. 33. Orecchia, P. et al. Electrophoretic identification of larvae and adults of Anisakis (Ascaridida:Anisakidae). J. Helminthol., 60, 331, 1986. 34. Orecchia, P. et al. Two new members in the Contracaecum osculatum complex (Nematoda, Ascaridoidea) from the Antarctic. Int. J. Parasitol., 24, 367, 1994. 35. D’Amelio, S. et al. Genetic variation of a population of the Pseudoterranova decipiens complex from Japan. Abstracts of the VIth European Multicolloquium of Parasitology, 7, 45. The Hague, The Netherlands, 1992. 36. Paggi, L. and Bullini, L. Molecular taxonomy in anisakids. Symposium on “Parasites of biological and economical significance in the aquatic environment”. Bull. Scand. Soc. Parasitol., 4, 25, 1994. 37. Arduino, P. et al. Isozyme variation and taxonomic rank of Contracaecum radiatum from the Antarctic Ocean (Nematoda, Ascaridoidea). Syst. Parasitol., 30, 1, 1995.

768 38. Farjallah, S. et al. Molecular characterization of larval Anisakid nematodes from marine fishes off the Moroccan and Mauritanian coasts. Parasitol. Int., 57, 430, 2008. 39. Umehara, A. et al. Multiplex PCR for the identification of Anisakis simplex sensu stricto, Anisakis pegreffii and the other anisakid nematodes. Parasitol. Int., 57, 49, 2008. 40. Abe, N. Application of the PCR-sequence-specific primers for the discrimination among larval Anisakis simplex complex. Parasitol. Res., 102, 1073, 2008. 41. D’Amelio, S. Phylogeny of anisakid nematodes: a review, Helminthologia, 40, 87, 2003. 42. D’Amelio, S. et al. Diagnosis of a case of gastric ­anisakidosis by PCR-based restriction fragment length polymorphism analysis. Parassitologia, 41, 591, 1999.

Molecular Detection of Foodborne Pathogens 43. Farjallah, S. et al. Diagnosis of a case of oesophageal anisakidosis by PCR-based restriction fragment length polymorphism analysis. Parassitologia, 49, 217, 2007. 44. Umehara, A. et al. Molecular identification of the etiological agents of the human anisakiasis in Japan. Parasitol. Int., 56, 211, 2007. 45. Mattiucci, S. et al. Human Anisakidosis in Italy: molecular and histological identification of two cases. Parassitologia, 49, 226, 2007. 46. Moneo, I. et al. Sensitization to the fish parasite Anisakis simplex: clinical and laboratory aspects. Parasitol Res., 101, 1051, 2007. 47. Caballero, M.L. and Moneo, I. Several allergens from Anisakis simplex are highly resistant to heat and pepsin treatments. Parasitol Res., 93, 248, 2004.

55 Clonorchis

Heinz Mehlhorn, Boris Müller, and Jürgen Schmidt Heinrich Heine University

Contents 55.1 Introduction.................................................................................................................................................................... 757 55.1.1 Classification and Morphology ........................................................................................................................ 757 55.1.2 Biology.............................................................................................................................................................. 759 55.1.3 Pathogenesis and Control.................................................................................................................................. 760 55.1.4 Diagnosis of Foodborne Clonorchiasis..............................................................................................................761 55.2 Methods...........................................................................................................................................................................761 55.2.1 Detection of C. sinensis rediae in Snails...........................................................................................................761 55.2.2 Detection of Metacercariae in Fish................................................................................................................... 762 55.2.3 Detection of C. sinensis Eggs in Human or Animal Fecal Samples................................................................ 763 55.3 Conclusions and Future Perspectives............................................................................................................................. 766 References.................................................................................................................................................................................. 767

55.1  Introduction For about 60% of mankind, fish account for a significant ­proportion (approximately 40%) of their protein intake. However, in recent decades, the world has witnessed a drastic decline in fish harvest from the oceans due to overfishing as well as to the increased incidences of fish diseases (e.g., Anisakis sp. and tapeworms) resulting from contamination of waterways and seas by human, agricultural, and industrial wastes. If eaten raw or undercooked—even in salted preparations, the infected fish, acting as intermediate hosts in the parasite’s life cycle, can cause severe diseases in humans and their pet animals (dogs, cats), who function as the final (definitive) hosts harboring the adult worms for these parasites.1–3 Therefore, establishment of feces-safe fish cultures, regular inspection of fish meat, proper cooking of deep frozen fish before consumption play essential roles in reducing potential fishborne diseases in human populations.

55.1.1 Classification and Morphology “The genus Clonorchis in the helminth group Digenea belongs to the trematodes (= sucking worms) within the phylum Platyhelminthes (flat worms). The name Digenea comes from Greek digenes = two sexes and refers to the fact that most of the Digenea are hermaphroditic organisms that combine several generations within their complicated life cycle. The genus name Clonorchis (Greek = klon, latin– ­clonus, Greek–opisthen = in the rear, orchis = testis, testicle) describes that the two deeply lobulated testes are situated behind the female system at the ­posterior region of the worm Figure 55.1). Clonorchis sinensis —the so-called Chinese liver fluke of humans—is one of the well  known parasites infecting humans.” Awareness of this

worm is old, since its eggs were already found in a corpse from the West Han Dynasty in China more than 2,300 years ago. However, the first valid observation of the adult worm was recorded 1875 by McConnell4 at the autopsy of a Chinese carpenter in Calcutta, India. The American scientist Cobbold1 used 1875 as genus name Distoma (= two mouth openings) referring to the two suckers. The German zoologist Looss was considered to have delivered in 1907 the first full and valid description of the C. sinensis life cycle,5 although the Japanese scientist Ijiima discovered 1887 that C. sinensis also occurs in some nonhuman hosts that live around humans and thus he established the concept of reservoir hosts, which make it difficult to eliminate this parasite from humans.6 Other Japanese—Kobayashi (1914) and Muto (1918)—contributed the confirmation of important stages in the life cycle showing that fresh water fish are the second intermediate host, while several water snails (e.g., members of the genera Bulimus, Bithynia) act as first intermediate hosts, within which the fish-infecting cercariae are produced.7,8 Recently the genus name Clonorchis is often replaced by Opisthorchis (Greek–opisthos = behind, orchis–testis), since two other species O. felineus and O. viverrini (see Chapter 60 of this book) are so closely related to C. sinensis that two different genus names (with the identical meaning) are not justified. Clonorchis (syn. Opisthorchis) sinensis is found focally in many countries of East Asia (Thailand, Laos, Cambodia, Korea, China, Taiwan, Hong Kong, Macao, Vietnam) with a decreasing tendency, although a true mass evaluation of its epidemiology had not been undertaken.9,10 Thus the number of infected humans is only based on estimations, which range between 9 and 30 million humans including thousands of Chinese immigrants in the USA. However, many more millions live under the risk of an infection, since people in 769

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Molecular Detection of Foodborne Pathogens

(a)

(b)

GÖ

V

(c)

BS

U V D O H H

LK

2mm

Figure 55.1  Adult worms of Clonorchis sinensis. (a) Light micrograph of a carmin-red stained C. sinensis worm (D = intestine, H =­testis, O = ovary, U = uterus filled with eggs, V = vitellarium). (b) Diagrammatic representation of an adult stage of C. sinensis (BS = ventral Ju-cker, GÖ = genital opening, LK = Laurer’s channel). (c) Scanning electron micrograph of an adult, contracted C. sinensis, showing the smooth surface and the two suckers, which in former times led to the genus description as “Distoma” = two mouths.

the endemic regions love it to eat in many variations raw or undercooked fish, which harbor metacercariae. When the potentially infected fish are transported to the large cities (e.g., from South China to Hong Kong), they pose health risk for a much larger population base. Adult C. sinensis worms show some characteristics which allow a clear and convincing species diagnosis: (1) Size. They are 10–28 mm in length and reach 3–7 mm in width with a thickness of about 1 mm. These variations are due to the fact that living worms may stretch and contract considerably by activity of their strong layers of circular and longitudinal muscles. (2) Shape. In the state of rest the adult worms appear slightly lanceolate or leaf-like (Figures 55.1 and 55.2) with a pointed/tapered anterior and a rounded posterior end. (3) Appearance. Living worms appear whitish translucent, so that the internal organs can be seen from outside without means of coloration. (4) Characteristics. Two suckers are present at the ventral side of the worms and are used to attach at the walls of the bile ducts. The anterior sucker surrounds the mouth and thus is called oral sucker, while the second one has no connection to any internal organ. This ventral sucker is situated just in a short distance behind the two genital pores at the end of the first third of the worm’s body. (5) Surface. The tegumental surface of the adults is not provided with hooks, which are found in

other trematodes such as Fasciola hepatica, some Schistosoma species in Metagonimus yokogawai or in its own juvenile stages. (6) Sex. The adult worms are hermaphrodites with two lobulated testes at the posterior end, being situated behind each other. The one in front has four main lobes, the one behind five lobes, which are further subdivided (Figure 55.1). From the testes the sperms are transported via one rather long vas efferens to a short common vas deferens which opens into the male sexual porus. At the end of the male porus a short penis-like cirrus can become protruded and introduced into the own or in case of a copulation— into a foreign female pore. The female system consists of several compartments: (i) a single ovarium (ovary, germarium); (ii) a single ootype (egg formation chamber) with an opening to the worm’s surface via the so-called Laurer’s channel; (iii) two vitellaria—which are glands being situated at each of both lateral sides of the worm. Both vitellaria produce supplementary cells, which are transported via a common channel into the ootype; (iv) a single receptaculum seminis, which is used as storage sack for sperms that arrive via the convoluted uterus, which has its starting point in the ootype and which transports the fertilized eggs by means of rhythmic constrictions to the female opening at the sexual porus; (v) several single cell glands (Mehlis’ glands) which surround the ootype and help with their products to fortify the egg shell which derives from excretions of the vitellarian cells.

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(7) Eggs. The oocyte—produced in the ovarium—is fertilized inside the ootype, the lumen of which has the shape of the later egg. The zygote becomes surrounded still inside the ootype by up/to 20 vitellarian cells which excrete the confluencing egg shell. This combination of zygote and helping vitellarian cells is mostly called “egg,” although its organization represents a cocoon. During the transport of the eggs/cocoons to the female sexual opening inside the convoluted uterus its wall becomes considerably hardened and finally appears yellowish brown with a length of 28–35 µm and a width of 12–19 µm Figure 55.2). They are characterized by an operculum (lid), at its anterior pole giving rise to a collar-like swelling. At the posterior end there is a knob-like swelling (dot). Inside the egg/cocoon the larval development starts already while it is still inside the uterus. The development of the larva—a motile miracidium—is finished when the egg has been excreted by the worm. The eggs leave the final host via the bile fluid and feces and must reach a convenient site (i.e., water). In contrast to other trematodes the miracidium of C. sinensis does not leave the egg shell in water, but waits until it has been engorged by a water snail. In the intestine of the latter, the operculum is blown up, the miracidium creeps out and enters the midgut gland of the snail (hepatopancreas) to proceed there its further development. (8) Intestine. The two trunks of the digestive system unite in a short esophagus just before the mouth and end blindly with no rectum or anus at their posterior ends. Thus all remnants of digested food have to become discharged via the mouth. (9) Excretion system. Watery excretion is done via socalled solenocytes = cyrtocytes = terminal cells of the protonephridial system. Their excretory channels unite in an enlarged channel, that ends in a large

bladder at the posterior end of the worm. From there the excretory fluids are released to the outside. (10) Nervous system. The nervous system is composed of an esophageal commissure with ladder-like pairs of nerve trunks running in each plane of the body until the posterior end of the worm. The nerves are provided with terminal sensory processes which are distributed along the surface of the integument (= neodermis). (11) Body cavity. There is no true body cavity in the interior of the worm below the layers of the syncytial integument and the annular and longitudinal muscles. This space is filled by interconnected parenchymal cells which fulfil transportation functions.

55.1.2  Biology The adult flukes—up to 20,000 specimens have been found within one patient—live within the bile ducts and feed on epithelia (Figure 55.2). Fully embryonated eggs are laid by the worms, pass from the common bile ducts into the small intestine and become excreted with the feces. From early animal experiments,11 it is concluded that each Clonorchis worm produces about 1,000–4,000 eggs per day. This rather limited production of eggs is probably the reason why infections with small numbers of adult Clonorchis will hardly be found by fecal examination techniques. If the eggs/ cocoons of Clonorchis had been passed into water, water snails of various genera (Bithynia, Bulimus, Semisulcospira, Parafossarulus, etc.) ingest such eggs. Inside their intestine the included larva hatches from the operculated egg and enters the midgut gland of the snail, where further development is initiated. The ciliated miracidium grows up into a sporocyst, within which rediae are produced. The latter produce a second generation of rediae, which finally give rise to motile cercariae. These stages consist of an anterior young worm with the typical anlagen of the later organs and a long

(a)

(b) OP

G M B

U

Figure 55.2  Light micrographs of Clonorchis sinensis. (a) Light micrograph of a section through Clonorchis worms in a human bile duct (B = ventral sucker, G = bile duct, U = uterus). (b) Light micrograph of a typical C. sinensis egg showing the typical apical operculum (OP) and the developing miracidium (M).

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(tail, which is up to four times longer than the young worm). This tail may propel the cercaria in water after its emerges from the snail. When getting close to a second intermediate host—one of more than 100 species of cyprinid or salmonid fish—the cercaria is attracted by skin odour12 and bores through the epidermis after discharging the tail. Having reached the dermis, the muscles or the subcutaneous tissues the small juvenile worm stages transform into metacercariae. Each of them excretes a smooth cover which protects the included worm from host cell attacks. Such metacercariae are spherical in shape with diameters of about 0.3 mm, appear whitish and thus are hardly seen with the naked eye in the whitish muscle tissue or when situated below the fish’s scales. As soon as these second intermediate hosts are eaten by humans, dogs, or cats (and other possible predator like swine when fed with food wastes), the young immature worms hatch from the metacercarian cover inside the small intestine and crawl up the bile duct and establish themselves there, growing to adulthood within 2 weeks. Generally C. sinensis locates within the small intrahepatic bile ducts. In heavy infections adult worms may also occur in the extrahepatic bile ducts and in the gall bladder.13 A tissue migration (as in young Fasciola hepatica worm) was never seen. The adults of C. sinensis may copulate or produce their eggs after self-fertilization, thus completing the life cycle. The life span of the adult worms may reach up to 25 years as it was claimed for schistosomes.14,15 However, in all cases with long lasting chlonorchiasis—Asians living far away from their native country—it could not definitively excluded that they had become reinfected, e.g., by eating salted raw fish. Thus summarizing the life cycle of C. sinensis inside man the following data were recorded in literature: (i) incubation period of about 14 days in heavy infections, otherwise without symptoms; (ii) prepatent period of at least 14 days; in low grade infections, however, eggs are often found much later; (iii) patency of up to 25 years.

55.1.3  Pathogenesis and Control Pathogenesis. The intensity of pathogenic effects of clonorchiasis depends on the amount of adults and pre-adults at their final sites of location: the bile ducts, which, however, influence the function of the whole gastrointestinal system. The basis of the damages seen in the bile ducts and gallbladder are inflammatory changes induced by the attachment of the two suckers at the wall of the parasitized ducts and their feeding behavior of engorging epithelial cells. The changes increase in number and in severity with the number of feeding adult worms and include desquamation of the biliary epithelium and formation of crypts. These effects result in a marked proliferation of the mucosa, a thickening of the walls (metaplasia) of the bile ducts and in a considerable dilation of the latter. Consequently there may be severe damage to the liver, which becomes enlarged in cases of acute disease. Such damages may lead to cirrhosis of the liver, oedema, and formation of ascites. Cancer of the liver and of the ducts were also seen, among them cholangiocarcinoma

Molecular Detection of Foodborne Pathogens

and hepatocarcinoma.2,10,14–22 In some cases even ­pancreatitis has been claimed to occur as consequence of an etiologic relation to severe infections with C. sinensis.23 The above described pathogenic effects of an infection with C. sinensis worms must not introduce well defined clinical symptoms. If the number of worms is rather low, e.g., in old chronic infections, only mild, nonspecific gastrointestinal symptoms occur, if at all. On the other hand severe infections are mostly associated with intensive right upper quandrant pain and tenderness. In any case an infection with C. sinensis must never be neglected, especially with respect to the long-time effects of a duct wall cancer, which has a very poor prognosis, since most of the patients died within 2 years after the diagnosis. The clinical symptoms of clonorchiasis thus should be grouped as follows: (i) First acute phase: this phase starts just 1 week after infection and may last for longer showing the following symptoms (singly or in combination): anorexia, feeling of stoutness, diarrhea, fevers, rigors, and chills, colic–like abdominal pain, slight icterus, enlargement of liver, weight loss, leucocytosis combined with eosinophilia. (ii) Chronic phase: with increasing persistence of the disease a subsequent chronic phase may show the following symptoms: gallbladder enlargement, edema, meteorism, cachexia, cholangitis, chronic diarrhea, increasing hepatomegalia, retardation of growth, and sexual maturity, and neuropsychiatric symptoms. (iii) Complications: as complications the following events may occur: ascending cholangitis, intermittent severe cholestasis, pancreatitis, liver cirrhosis, formation of gallbladder stones (hepatolithiasis), obstructive jaundice, ascites, portal hypertension, secondary bacterial infections, night-blindness due to lack of vitamin A, cholangionergic liver carcinoma gastrointestinal bleedings, death due to multi-organ failure. Prophylaxis. With respect to the possibility of severe symptoms of disease as follow up of an infection with C. sinensis, measurements of prophylaxis must be undertaken. At first it is needed to create awareness in the population on the dangerous custom of eating raw or undercooked fish. The second need is to cook or boil any fish, at least those from fresh or brackish water, since the latter might have been contaminated by worm-egg-containing human or animal feces and thus are a source of infection of fish with these trematodes. Another possibility of prophylaxis is shock frosting of freshly caught fish. This procedure can be used in industrial plants and would avoid foodborne infections, even if later the fish are eaten raw. Finally the implementation of a feces collecting system in villages would exclude the import of possibly worm egg-contaminated feces into fish ponds. Therapy. Fortunately the chemotherapy of clonorchiasis and related worm diseases is rather uncomplicated due to the discovery of the compound praziquantel, which is the drug of choice against trematodes and cestodes since the year

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1979 until now. This drug is generally well tolerated and has a very high efficacy. Thus there are up to now no proven cases of resistance. Praziquantel enters the blood soon after its oral uptake and thus it reaches the tissues within a few minutes.24 The reported side effects (malaise, headache, dizziness) remain rather mild. Even the occasionally occurring symptoms like sedation, abdominal discomfort, slight fever, sweating, nausea, eosinophilia or fatigue are not very important as are not the rarely found pruritus or rash. It was shown that 3 × 25 mg praziquantel per kg bodyweight on each of two successive days or 20 mg praziquantel per kg body weight for 3 days are sufficient for a complete cure in cases of clonorchiasis and opisthorchiasis.24–27

55.1.4  Diagnosis of Foodborne Clonorchiasis Diagnosis of clonorchiasis has relied for long exclusively on the demonstration of the very small and tiny eggs of this worm by methods of stool examination like merthiolate-­iodineformaldehyde concentration (MIFC) or sodiumacetate-acetic acid formaline (SAF). These methods used worldwide are not very specific, however, they give a rather good general survey on many parasites which may occur in human or animal feces. In cases of mild infections with a low number of Clonorchis worms, the number of the daily excreted eggs is also low so that they may be easily overlooked. Furthermore there is obviously no possibility of definite discrimination between Clonorchis and the eggs of the species Opisthorchis felineus and O. viverrini. The same is true when the flotation method (zinc chloride/sodium chloride) is used.2 Adult worms can also be detected in the bile ducts by ultrasonification or by radiographical techniques, by endoscopic retrograde cholangiopancreatography. The different serological methods of ELISA, when first established 1992 by Feldheim and Knobloch,28 turned out to be very helpful, but soon cross reactions with the antigens of other genera of trematodes (e.g., Fasciola, Paragonimus, Schistosoma) were seen.26 Thus without use of molecular biological methods, which, however, are costful, and need moore time, a clear diagnosis of Clonorchis sinensis at the species level remains always somewhat doubtful. A similar situation is given with respect to the metacercariae inside the intermediate host fish. When practising the regular meat inspection of fish fillets, the metacercariae are very easily overlooked due to their small size of often less than 1 mm, by their whitish color in whitish fillets, by their small number or by their often hidden position below the scales. Furthermore genus and species determination remains rather doubtful using this time-wasting method, even if the digestion technique (used in trichellinosis) is added. Thus the use of molecular biological methods is required in order to detect an infection with C. sinensis.

55.2  Methods In this chapter, we present procedures for molecular detection of C. sinensis rediae in snails, metacercariae in fish, and

eggs in human or animal fecal samples, along with relevant sample collection and preparation techniques.

55.2.1  Detection of C. sinensis rediae in Snails The following detection method for C. sinensis sporocysts and rediae in snails has the advantage that very few steps have to be carried out, and that only inexpensive chemicals are required. Sample collection and preparation. Each snail is collected in a 2-ml Eppendorf tube, and the shell is crushed by using clean sticks. An amount of 1.5 ml of 25 mM potassium hydroxide is added to the tube and the snail’s soft body is lysed by incubation at about 95°C for 15 min. After a brief centrifugation to sediment the shell particles, about 10 µl of a saturated aqueous solution, cresol red, is added, and the supernatant is neutralized by the addition of 1 N HCl. The indicator dye appears violet at alkaline pH and becomes yellow when an appropriate pH ≤ 7.5 is reached. Of this sample, 1 µl is added to a PCR assay volume of 50 µl. The muscular food of the snail may not be entirely lysed during the short incubation. However, the rediae reside in the hepatopancreas and are easily freed from this soft tissue. The rediae are completely lysed by alkaline treatment, and the parasite DNA is efficiently liberated. It has been amply evaluated29 that it is not necessary to apply methods for further purification of the DNA, e.g., by using commercial kits. PCR. A suitable primer pair for the specific detection of C. sinensis has been designed from the internal transcribed spacer 2 (ITS2) of the ribosomal DNA after examination of the sequences of several related opisthorchiid species.29 The primer pair consists of CS1 (5´-CGAGGGTCGGCTT ATAAAC-3′) as a forward primer and CS2 (5´-GGAAAGT TAAGCACCGACC-3′) as a reverse primer. These primers yield a single DNA fragment of a typical product size of 315 bp. The utility of this PCR for the detection of rediae in snails or metacercariae in fishes was previously assessed with adult C. sinensis (stored in 70% ethanol, originating from Korea, Prof. Dr. Rim) as source material, with DNA extracted from the liver flukes with the commercially available QIAamp DNA mini kit (Qiagen, Hilden, Germany). The specificity of the PCR primers was examined with DNA from the closely related liver flukes Opisthorchis viverrini, O. felineus, Pseudoamphistomum truncatum, and Metorchis xanthosomus. In addition, DNA from the Digenea Echinostoma caproni, Fasciola hepatica, Schistosoma mansoni, Holostephanus dubenini, and Paracoenogonimus ovatus were tested as the DNA from intermediate hosts snails B. fuchsiana and Melanoides tuberculata and the cyprinid fishes P. parva, and Carassius auratus auratus, respectively.29 The PCR assay is carried out with a final volume of 50 µl with 1 × PCR buffer containing Tris-HCl, KCl, (NH4)2SO4 and 1.5 mM MgCl2 pH 8.7 (Qiagen), 1.25 U Taq DNA polymerase (Qiagen), 200 µM of each dNTP, 0.4 µM of each CS primer, and 1 µl of snail lysate (template DNA). PCR amplification is performed in a PTC-0150 MiniCycler (MJ Research, Watertown, MA) with the following cycling

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conditions: one cycle of 94°C for 3 min; 40 cycles of 94°C for 1 min, 62°C for 1 min and 72°C for 1 min; and one cycle of 72°C for 10 min. After the completion of the cycles, the amplified PCR products of 315 bp in size are separated in agarose gel and visualized with ethidium bromide stain. Note. PCR has been successful performed with snail lysate from Bithynia fuchsiana infected with C. sinensis rediae, as the substances inhibiting the enzymatic reaction are eliminated in the lysate preparation. This was confirmed by adding 1 µl of lysate from uninfected snails to the PCR assay, together with serial diluted genomic DNA from C. sinensis. The PCR assay based on the primer pair CS1/CS2 has a detection limit of 10 –13 g DNA. Clearly, the sample processing technique has successfully removed interfering components from the snail tissue as an identical level of sensitivity is achieved with parasite DNA tested alone or in the presence of snail lysate (Figure 55.3). Using a preparation volume of 1.5 ml, the PCR is capable of detecting 1.5 × 10 –10 g DNA of rediae from an infected snail. With this level of sensitivity, practically any infected snail will be recognized as positive (a)

M

1

2

3

by the PCR assay. Thus, the PCR procedure provides a ­reliable means for identifying C. sinensis-infected snails.

55.2.2  Detection of Metacercariae in Fish Traditionally, metacercariae isolated from fish are detected by light microscopy. Given that all mentioned digenean metacercariae form ovoid cysts and have thick-walled, ovoid excretory vesicles filled with dark corpuscles, it is difficult to differentiate umambiguosly metacercariae of C. sinensis from those of related human pathogenic species of the genera Opisthorchis and Heterophyes and also from other less known and less pathogenic digenean species. To determine the specimens reliably, it may be necessary to infect susceptible mammals with the metacercariae to obtain adult worms. This is unduly cumbersome and time-consuming. The following method allows detection of C. sinensis metacercariae in fish without microscopical inspection:29 Sample preparation. Fish filets (2–3 g) are taken from Pseudorasbora parva fish, finally minced, and 4

5

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Figure 55.3.  (a) Sensitivity of the CS1/CS2 primers with purified C. sinensis DNA. Lane M, DNA size marker; lane 1, 10–10 g; lane 2, 10–11 g; lane 3, 10–12 g; lane 4, 10–13 g; lane 5, 10–14 g; lane 6, 10–15 g; lane 7, 10–16 g; lane 8, 10–17 g of C. sinensis DNA; lane 9, negative control. (b) Detection of C. sinensis metacercariae from fish with the CS1/CS2 primers. One metacercaria was artificially inoculated into each 3 g fish meat prior to processing the samples. Lane M, DNA size marker; lanes 1–10, samples from infected fish; lane 11, negative control containing no metacercaria. (c) Detection of C. sinensis eggs in human stool with OP1/OP2 primers. One egg was inoculated into each 0.1 g stool sample before processing. Lane M, 50-bp ladder; lanes 1–10, stool samples; lane 11 negative control containing no egg.

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then transferred to a 50-ml Falcon tube containing artificial ­gastric juice ­consisting of 0.9% pepsin, 0.7% HCL, and 1% Triton X-100. Use of detergent Triton X-100 here improves the dissolution of the fish muscles and does not destroy the metacercarial cysts. This mixture is incubated at 38–40°C and occasionally shaken by hand. Digestion is complete within 45–60 min. Then, the samples are briefly centrifuged to sediment the metacercariae. The sediment is washed twice by removing the supernatant and filling it up with water. Subsequently, the supernatant is removed but left a residual volume of 1–1.5 ml. The disruption of the metacercariae and the liberation of the DNA are achieved by homogenization with a tissue homogenizer (Omni TH220, Omni International, Marietta, GA) at 10,000 rpm for 1 min. After a brief centrifugation, the supernatant is boiled in a water bath for 5 min to inactivate the pepsin and, subsequently, is neutralized by the addition of 1 N potassium hydroxide. Neutralization is carried out with the aid of cresol red as indicator dye. Cresol red appears yellow at acid pH and becomes violet when an appropriate pH of around ≥ 7.5 is reached. 2 µl of this sample is added to 50 µl PCR assay volume. This preparation procedure for metacercariae from fishes is rapid and simple as it is not involve further extraction of DNA by phenol-chloroform or even more complex purification methods. Inhibitors contained in fish samples are effectively removed by simple washing steps, whereby they are diluted to a value that enables the PCR to reliably work. PCR. This is conducted with the CS primers as outlined in Section 55.2.1 Detection of C. sinensis rediae in snails, with the positive specimens showing an expected single DNA fragment of 315 bp. Note. Reports on molecular methods to detect opisthor­ chiid liver flukes are scarce. To our knowledge for C. sinensis, only one study with PCR has been published,30 in which a mitochondrial-based multiplex PCR for the identification of C. sinensis and O. viverrini was described, with a sensitivity of 7.8 × 10 –10 g of genomic parasite DNA. The CS primer pair developed for C. sinensis by Müller et al.31 is, however, much more sensitive with a detection limit of 1 × 10 –13 g DNA. By evaluating with multiple samples, the reliability of the DNA preparation procedure for metacercariae from fish is validated. To do this, fresh fish fillet from ten uninfested P. parva, bred parasite-free in the laboratory,29 were inoculated with one metacercariae of C. sinensis (Figure 55.3). Examination of DNA from closely related opisthorchiid liver fluke species, other Digenea and various intermediate hosts mentioned above showed no cross-reactions with the primers specific for C. sinensis. These results indicate that the PCR-based method is a suitable tool for the detection of C. sinensis in infected fish, with sensitivity approaching to only one metacercariae per fish sample. In contrast to conventional techniques, PCR-based methods have a higher potential for the differentiation of digenean species, which is essential for epidemiological studies. Thus, application of the PCR enables an unambiguous identification of C. sinensis, which

can be difficult by light microscopy. This technique offers a better way to track the relevant pathways of transmissions by identifying host species and infection reservoirs. With respect to the expanding worldwide trade of fresh water fish from aquacultures, it is highly desirable to have a means for control inspections for fish in order to avoid importation of fish being infected with C. sinensis metacercariae.32

55.2.3 Detection of C. sinensis Eggs in Human or Animal Fecal Samples The opisthorchiids live in the bile ducts and the gall bladder (Figure 55.2). All species cause similar symptoms, such as adenomatous hyperplasia of the biliary epithelium or obstruction of the biliary system. Chronic infections also may introduce liver cirrhosis, cholangiocarcinoma, or liver carcinoma.9,33 It appears that species of the genera Metorchis and Pseudamphistomum occasionally infect humans and cause the same problems as the better known liver flukes.34 Since all species of the opisthorchiids cause the same health problems it makes sense to use a single test system for their diagnosis. However, for appropriate treatment of patients, e.g., with praziquantel,24,26 there is no need to determine the exact species. The development of PCR techniques permits identification of flukes of the family Opisthorchiidae from human and animal fecal specimens, and discrimination of all opisthorchiids from other species of Digenea, especially from the members of the related family Heterophyidae or other so-called minute flukes that colonize in the intestine of humans. Cleanup procedure. It is well-known that feces contain various substances that strongly inhibit amplification by PCR.35 The conventional methods for extraction of DNA of parasite eggs from stool samples are complex, and require use of expensive commercial kits to remove inhibitory substances. Many of these DNA preparation procedures are based on those for the diagnosis of bacteria and viruses, and do not take account of the fact that DNA of the parasite is enclosed by the egg shell. Not surprisingly, the DNA so obtained does not always give reproducible PCR amplification as the DNA polymerase inhibitors are not completely eliminated from fecal material. The present technique can be carried out rapidly with inexpensive chemicals, and even low amounts of worm eggs in feces can be detected reliably by PCR. As reported by Müller et al.31 100 mg of fecal samples are collected in a 2-ml Eppendorf tube and incubated with an aqueous solution consisting of 2% Triton X-100 and 25 mM potassium hydroxide for 10 min at room temperature on a shaker. After centrifugation at 10,000 rpm in an Eppendorf centrifuge for 2 min, the supernatant is removed, and the sediment is washed two or three times with 2% Triton X-100 and 25 mM KOH solution until the supernatant appears clear. Then, the supernatant is discarded and 0.8 ml of sucrose/ potassium chloride solution is added to the sediment containing the eggs. The sucrose/KCl solution consists of 82.5 g sucrose, 20 g KCl, and distilled water added up to 100 ml.

776

This solution has a density of 1.39 g/ml. During centrifugation at 14,000 rpm for 5 min, the eggs are distributed within the sucrose/KCl solution and do not float up to the surface of the solution. Thus, they are separated from the sedimenting fecal particles. The entire supernatant containing the eggs is transferred into a new 2-ml tube, which is then filled up with water and centrifuged at 14,000 rpm for 5 min. By adding water, the density of the solution is lowered so that the eggs became sedimented. The supernatant is removed and the sediment washed twice with water. After the second washing, the supernatant is removed, leaving a remnant of 0.3 ml. The disruption of the egg shell and release of the parasitic DNA are achieved by homogenization of the sample with a tissue homogenizer (Omni TH220, Omni International, USA) at 10,000 rpm for 2 min. Alternatively, the eggs are opened by autoclaving.36 However, the autoclaving approach is found to be less efficient since a considerable percentage (often 50%) of the eggs remain closed. After a brief centrifugation, the supernatant is boiled in a water bath for 5 min to inactivate hydrolytic enzymes of the feces. 5 µl of this sample, free of inhibitory substances, is added to the PCR tube in a final volume of 50 µl. PCR. The species of the family Opisthorchiidae share extreme homologies of the internal transcribed spacer 2 (ITS2) sequence Figure 55.4), which is clearly different from that of other Digenea. Thus, this sequence is exploited for development of specific primers OP1/OP2 for the opisthorchiid species C. sinensis, Opisthorchis viverrini, Opisthorchis felineus, Pseudamphistomum truncatum, and Metorchis xanthosomus. Indeed, C. sinensis sequences derived from specimens from Korea (AF217094) and from China (AF217099) demonstrate identical compositions. Sequences from O. viverrini (AF408147) are available in GenBank. Data for O. ­viverrrini that had been reported for isolates from different areas in northeast Thailand37 are also identical with one another and with those shown in GenBank. Nucleotide sequences of the ITS2 region of O. felineus, P. truncatum, and M. xanthosomus have not been reported before and are determined (Sequence Laboratories Göttingen GmbH, Germany) (Figure 55.4). Although DNA from F. hepatica generates an amplification product, this product is not in the range of the specific products of the opisthorchiid flukes. The sensitivity of the primers is determined in assays with diluted genomic DNA of C. sinensis, and a minimal amount of 10 –15 g DNA is achievable. The primer pair consists of the forward primer OP1 (5′-CGAGGGTCGGCTTATAAAC-3′) and the reverse primer OP2 (5′-AGCCTCAACCAAAGACAAAG-3′), yielding a specific 250 bp product from the family Opisthorchiidae. The PCR is carried out in a final volume of 50 µl, consisting of 1 × PCR buffer [Tris-HCI, KCI, (NH4) 2 SO4, 1.5 mM MgCI2; pH 8.7], 1.25 U Taq DNA polymerase (Qiagen), 200 µM of each dNTP, 0.4 µM of each primer (OP1 and OP2) and 5 µl DNA. PCR amplification is conducted in a PTC0150 Minicycler (MJ Research, USA) with the following cycling conditions: one cycle of 94°C for 3 min; 40 cycles of

Molecular Detection of Foodborne Pathogens

94°C for 1 min, 62°C for 1 min, and 72°C for 1 min, and one cycle of 72°C for 10 min. The amplified products of 250 bp in size are separated by agarose gels and stained with ethidium bromide. Note. The cleaning up procedure and efficacy of the PCR have been evaluated with an array of stool samples to confirm their reliability and extremely high sensitivity. DNA from adult C. sinensis originating from Korea was used for the establishment of the PCR assay. The DNA extraction of the flukes is also carried out with the commercially available QIAamp DNA Mini Kit (Qiagen, Hilden, Germany). The PCR yielded amplification products of approximately 250 bp for all tested opisthorchiid flukes, i.e., C. sinensis, O. viverrini, O. felineus, P. truncatum, and M. xanthosomus. To evaluate possible cross-reactions with DNA from other Digeneans, Echinostoma caproni, Fasciola hepatica, Schistosoma mansoni, Holostephanus dubenini, and Paracoenogonimus ovatus are analyzed. DNA from several intermediate hosts, the snails Bithynia tentaculata, Bithynia leachi, Bithynia fuchsiana, Bithynia funiculata, and Melanoides tuberculata and the cyprinid fishes Pseudorasbora parva, Rutilus rutilus, and Carassius auratus, are also tested. DNA from other Digenea and from intermediate hosts does not cross-react with the OP1 and OP2 primer pair. Additionally, the PCR using OP1/OP2 primers is run with each of ten 0.1-g stool samples of uninfested humans experimentally spiked with only one Clonorchis sinensis egg (Figure 55.3). All samples give a strong signal in PCR. For convenience, the tests can be carried out using Eppendorf tubes containing 0.1 g feces dotted with the lowest number, i.e., one egg. Hence, the proven sensitivity of the method can be calculated as 10 eggs per gram (EPG) feces. However, if diagnosis is carried out with 1-g stool samples in larger test tubes, the sensitivity may be estimated to be under 10 EPG. The sample preparation method has originally been developed using feces of cats infected with C. sinensis. It is later found to work comparably well using either human or cat feces. The purification method, however, is less suitable for feces of rodents containing large amounts of undigested plant materials. The technique can also be used for species- or strain­specific diagnosis of other parasites, such as eggs of nematodes or cestodes, oocysts of coccidian species (Apicomplexa), cysts of amoebae and of Giardia species, or of other protozoan species occurring in feces. Diagnosis on conventional light microscopy is difficult in patients with low infections, who shed less than about 200 EPG feces. Methods like Stoll’s egg count and formalin-ether technique easily fail in cases of low infection rates.13,36 The sensitivity of light microscopy diagnosis for O. viverrini egg in stool samples was examined in comparison with numbers of worms recovered from the liver of autopsied humans.13 According to this study, only in 30% of patients harboring one to nine worms (22% of all infected persons) did the stool diagnosis give a positive result. In cases of burdens with ten to 19 worms (11% of infected humans), the result was positive

Clonorchis

777

Figure 55.4  Nucleotide sequences of the ITS2 region of opisthorchiid flukes. Nucleotide sequences of O. felineus, P. truncatum, and M. xanthosomus were determined. The asterisks show those base pairs that are identical in all five species. The sequences of C. sinensis (AF217094/AF217099) and O. viverrini (AF408147) are registered in GenBank.

only in 77% with the Stoll’s and in 61.5% with the formalinether technique. The eggs of opisthorchiids are peculiar because in flotation media, low numbers of eggs are hardly detectable. Harnnoi et al. compared the flotation of O. viverrini eggs in saturated sodium nitrate solution (specific gravity 1.4) to the results when using the formalin-ether sedimentation. Attempts to

float eggs quantitatively failed; thus, the authors revealed that the flotation technique was not as efficient as the sedimentation. The specific gravity of O. viverrini eggs was found to range from 1.27 to 1.30 g/ml.38 Although the specific gravity of O. viverrini eggs is lower than that of the salt solution (specific gravity 1.4), the eggs do not float. An additional flotation technique is the zinc chloride/sodium chloride method.2

778

The specific gravity of this solution is 1.3 and failed, as the attempt with saturated sodium nitrate solution (specific gravity 1.4), to float the eggs quantitatively. Thus, it appears that flotation techniques are not very suitable for the detection of opisthorchiid eggs in feces in cases of low infection rates. The egg production of a single C. sinensis worm has been determined to be 100 EPG of feces.11 Song et al.26 reported in their 1986 study on the mean EPG of C. sinensis patients in various areas in Korea that the lowest single value found was 100 EPG. Haswell-Elkins et al.10 and Sithithaworn et al.13 found that one O. viverrini worm releases about 50 to 200 EPG host feces in 1991. With the present sample preparation technique and PCR, one egg per 100 mg sample gave a strong signal. Calculating for a 1-g sample, the detection limit would be 10 EPG, and an infection with only a single worm is reliably detectable. Thus, the PCR based method, along with the reported cleanup procedure reported herein, is more sensitive and reliable than the normal light microscopy methods. Eggs of closely related opisthorchiid and heterophyid species morphologically appear very similar. A differentiation of the eggs of C. sinensis, O. viverrini, or O. felineus by light microscopy requires considerable skill. The small eggs of these species are ovoid in form, similar in shape and possess an operculum with a prominent shoulder (Figure 55.2). The present cleanup procedure for stool samples and the application of the PCR system OP1/OP2 is a robust and highly sensitive method to detect infections with opisthorchiid liver flukes. Thus, the reported techniques represent a suitable diagnostic tool. By employing the cleanup procedure reported here for stool samples together with a PCR using species-specific primers, it is possible to rapidly and reliably determine the species identity of opisthorchiids, which is important for epidemiological studies. For example, the use of the primer pair CS1/CS2 is recommended for species-specific detection of C. sinensis.29,31

55.3 Conclusions and Future Perspectives Clonorchis sinensis is parasitic trematode whose eggs readily enter suitable snails (as the primary intermediate host) to develop into rediae, which in turn infect fish (as the secondary intermediate host) and develop into metacercariae. When infected fish harboring metacercariae are consumed raw or undercooked by humans and animals (as final hosts), the metacercariae mature into adult worms whose eggs are excreted with the feces. Given that fish make up a significant source of dietary proteins for many countries, clonorchiasis has been an important foodborne disease in regions such as East Asia where raw fish are often consumed. Traditionally, laboratory diagnosis of clonorchiasis is dependent on the identification of C. sinensis eggs in the fecal samples, and other stages (rediae, cercariae, and metacercariae) in the intermediate hosts such as snails and fish. Since C. sinensis adult worms produce a limited number of

Molecular Detection of Foodborne Pathogens

eggs per worm, human or animal hosts harboring relatively few worms are not easily diagnosed with fecal examination techniques. The fact that C. sinensis rediae and metacercariae are small and share similar morphological features with those from other pathogenic species of Opisthorchis and Heterophyes genera makes the microscopic differentiation difficult. Given that clonorchiasis can become a severe disease in humans and animals, it is vital that a diagnosis even of a low worm burden is achieved as early as possible. The application of biochemical and serological techniques has enhanced the laboratory identification of C. sinensis to a large extent, but it only became possible to rapidly and precisely distinguish C. sinensis eggs, rediae and metacercariae with the recent development of PCR-based nucleic acid detection assays. These molecular techniques are also valuable for detecting small amounts of C. sinensis eggs that may be easily overlooked in feces with standard microscopical techniques. In particular, the use of OP1 and OP2 primers derived from the internal transcribed spacer 2 (ITS2) of the family Opisthorchiidae facilitates separation of eggs of opisthorchiid species from those of heterophyid species. Coupled with an efficient cleanup procedure, the PCR using the OP1/OP2 primers offers a quick and sensitive screening system for infections with opisthorchiid liver flukes. Furthermore, the application of another primer pair (CS1/CS2) from the internal transcribed spacer 2 (ITS2) of the ribosomal DNA of C. sinensis enables species-specific identification of C. sinensis rediae and metacercariae.29,31 There is no doubt that future optimization of a real-time PCR detection platform will further streamline the laboratory diagnosis of C. sinensis and contribute to the improved control and prevention of foodborne clonorchiasis in humans worldwide. Utilizing the current state of technology, diagnosis in human patients with opisthorchiids should be carried out best with the opisthorchiid-specific primer pair OP1/OP2. Assay with these primers are very robust and sensitive, since only 10 –15 g DNA can be detected. It is no problem to diagnose even the lightest infections (with one self-fertilizing worm) in humans; if there is an egg in a sample, the assay will detect it. The general use of this primer pair should be promoted. The three opisthorchiid species O. viverrini, O. sinenesis, and O. felineus, are astonishingly closely related. Pathology and treatment are identical. Hence, it is NOT necessary to determine the exact species in medical diagnosis since treatment is identical. For scientific purposes, (C.) O. sinensis-specific primer pair CS1/CS2 can be utilized, which enables a highly sensitive detection with fecal material also. The detection limit with CS1/CS2 is 10 –13 g parasite DNA in a single assay. This level of sensitivity easily surpasses the performance of other primers reported in the literature using a nested PCR format. For practical and economic reasons, the nested PCR should be avoided whenever possible. Similarly, despite its convenience and speed, real-time PCR is much too expensive at the moment and, thus, is unsuitable as a widely applied routine diagnostic tool, especially in developing countries.

Clonorchis

References



1. Despommier, D.D., Gwadz, R., and Hotez, P.J. Parasitic Diseases. Springer Verlag, Heidelberg, 1995. 2. Mehlhorn, H. et al. Diagnosis and treatment of parasitic of man. 2. circulation, pp. 6–7. Gustav Fischer Verlag Stuttgart, Jena, New York, 1995. 3. Mehlhorn, H. Encyclopedia of Parasitology, Vol. 1–2. Springer Heidelberg, New York, 2008. 4. McConnell, J.F.P. Anatomy and pathological relations of a new species of liver fluke. Lancet, 2, 271, 1875 5. Loos, A. Clonorchis (1907) Cited in: Yamaguti, S. Synopsis of Digenetic Trematodes of Vertebrates. Keigaku, Tokyo, 1971. 6. Ijiima, I. Notes on Distoma endemicum Baelz. J. Coll. Sci. Imperial Univ. Jpn., 1, 47, 1887. 7. Kobayashi, H. (1914). Fish as intermediate hosts in clonorchiasis. Cited in: Despommier, D.D., Gwadz, R., and Hotez, P.J. Parasitic Diseases. 34, Springer Verlag, Heidelberg, 1995. 8. Muto, S. On the primary intermediate host of Clonorchis ­sinensis. Chuo Igakakai Zasshi, 25, 49, 1918. 9. Song, I.C., Lee, J.S. and Rim, H.J. Epidemiological studies on the distribution of Clonorchis sinensis infection in Korea (in Korean). Korean Univ. Med. J., 20, 165, 1983. 10. Haswell-Elkins, M.R. Lung and liver flukes. In: Cox, F.E.G. et al. (eds). Parasitology, Topley and Wilson’s Microbiology and Microbial Infections, 10th ed. pp. 508–519, Hodder Arnold, ASM Press, Washington, DC, 2005. 11. Wykoff, D.E. Studies on Clonorchis sinensis IV: Production of eggs in experimentally infected rabbits. J. Parasitol., 45, 91, 1959. 12. Haas, W., Granzer, M., and Brockelman, C.R. Opisthorchis viverrini: finding and recognition of the fish host by the cercariae. Exp. Parasitol., 71, 422, 1990. 13. Sithithaworn, P. et al. Relationship between faecal egg count and worm burden of Opisthorchis viverrini in human autopsy cases. Parasitology, 102, 277, 1991. 14. Piekarski, G. Medizinische Parasitologie in Tafeln. Springer Verlag, Heidelberg, 1987. 15. Chai, J.Y., Murrel, K.D. and Lymbery, A.J. Fish-borne parasitic zoonoses: status and issues. Int. J. Parasitol., 35, 1233, 2005. 16. Kim, Y.H. Carcinoma of the gallbladder associated with clonorchiasis: clinicopathologic and CT evaluation. Abdom. Imaging, 28, 83, 2003. 17. Leung, J.W. et al. Hepatic clonorchiasis – a study by endoscopic retrograde cholangiopancreatography. Gastrointest. Endosc., 35, 226, 1989. 18. Leung, J.W. and Yu, A.S. Hepatolithiasis and biliary parasites. Bull. Clin. Gastroenterol., 11, 681, 1997. 19. Chen, M.G., Lu, Y. and Hua, X.J. Progress in assessment of morbidity due to Clonorchus sinensis infection: a review of recent literature. Trop. Dis. Bull., 91, 7, 1994. 20. Bunnag, D. and Harinasuta, T. Opisthorchiasis, clonorchiasis and paragonimiasis. In: McGraw, R.P., and Mclvor, D. (eds). Tropical and Geographic Medicine. pp. 461–469, McGrawHill, New York, 1984.

779 21. MacFadzean, A.J.S. and Yeung, R.T.T. Acute pancreatitis due to Clonorchis sinensis. Trans. R. Soc. Trop. Med. Hyg., 60, 466, 1966. 22. Hou, P.C. The relationship between primary carcinoma of the liver and infestation with Clonorchis sinensis. J. Pathol. Bacteriol., 72, 239, 1965. 23. Komiya, Y. Clonorchis and clonorchiasis. Adv. Parasitol., 4, 53, 1966. 24. Mehlhorn, H. et al. Ultrastructural investigations on the effects of praziquantel on human trematodes from Asia (Clonorchis sinensis, Metagonimus yokogawai, Opisthorchis viverrini, Paragonimus westermani, Schistosoma japonicum). Drug Res., 33, 91, 1983. 25. Rim, H.J. et al. Clinical evaluation of the therapeutic efficacy of praziquantel (Embay 8440) against Clonorchis sinensis infection in man. Ann. Trop. Med. Parasit., 75, 27, 1981. 26. Rim, H.J. The current pathology and chemotherapy of clonorchiasis. Korean J. Parasitol., 24 (Suppl. 3), 27, 1986. 27. Hänel, H. and Raether, W. Chemotherapy. In: Mehlhorn, H. (ed). Encyclopedia of Parasitology, Vol. 1–2. Springer, Heidelberg, 2008. 28. Feldheim, W. and Knobloch, J. Serodiagnosis of Opisthorchis viverrini infestation by an enzyme immuno-assay. Tropenmed. Parasitol., 33, 8, 1982. 29. Müller, B., Schmidt, J. and Mehlhorn, H. Sensitive and species – specific detection of Clonorchis sinensis by PCR in infected snails and fishes. Parasitol. Res., 100, 911, 2007. 30. Le, T.H. et al. Clonorchis sinensis and Opisthorchis viverrini: development of a mitochondrial-based, multiplex PCR for their identification and discrimination. Exp. Parasitol., 112, 109, 2006. 31. Müller, B., Schmidt, J. and Mehlhorn, H. PCR diagnosis of infections with different species of Opisthorchiidae using a rapid clean-up procedure for stool samples and specific primers. Parasitol. Res., 100: 905, 2007. 32. WHO, Manual of basic techniques, Geneva, Switzerland, 1995. 33. Schwartz, D.A. Review: helminths in the induction of cancer: Opisthorchis viverrini, Clonorchis sinensis and cholangiocarcinoma. Trop. Geogr. Med., 32: 95, 1980. 34. Schuster, R. and Leberegelbefall, In Aspöck H., Amoeben, Bandwürmer etc. und parasitäre Erkrankungen des Menschen in Mitteleuropa. Denisia, 6, 291, 2002. 35. Wilde, J., Eiden, J. and Yoken, R. Removal of inhibitory substances from human fecal specimens for detection of group A rotaviruses by transcriptase and polymerase chain reactions. J. Clin. Microbiol., 28, 1300, 1990. 36. Wonggratanacheewin, S. et al. Detection of Opisthorchis viverrini in human stool specimens by PCR. J. Clin. Microbiol., 40, 3879, 2002. 37. Ando, K. et al. Nucleotide sequence of mitochondrial CO I and ribosomal ITS II genes of Opisthorchis viverrini in northeast Thailand. Southeast Asian J. Trop. Med. Public Health, 32 (Suppl 2), 17, 2001. 38. Harnnoi, T. et al. Specific gravity of Opisthorichis viverrini egg. J. Helminthol., 72, 359, 1998.

56 Diphyllobothrium

Jean Dupouy-Camet and Hélène Yera Paris Descartes University

Contents 56.1 Introduction.................................................................................................................................................................... 781 56.1.1 Classification and Morphology......................................................................................................................... 781 56.1.2 Biology and Pathogenesis................................................................................................................................. 781 56.1.3 Conventional Diagnostic Techniques................................................................................................................ 782 56.1.4 Molecular Diagnostic Techniques.................................................................................................................... 783 56.2 Methods ......................................................................................................................................................................... 785 56.2.1 Reagents and Equipments................................................................................................................................. 785 56.2.2 Sample Preparation .......................................................................................................................................... 785 56.2.3 Detection Procedure......................................................................................................................................... 785 56.3 Conclusions and Future Perspectives............................................................................................................................. 786 References.................................................................................................................................................................................. 787

56.1  Introduction Tapeworms of the genus Diphyllobothrium (Cobbold, 1858) are widely distributed all around the world, with some being agents of human diphyllobothriasis, one of the most important fishborne zoonosis caused by a cestode parasite. Piscivorous birds and mammals are definitive hosts. The intermediate hosts include both freshwater and marine fish, mainly anadromous. Humans can contract this parasite by eating raw or partially cooked fish containing Diphyllobothrium plerocercoids. This cestode is commonly present in the wildlife among fish, mammal, and bird hosts that make an important reservoir.1–3 In 1999, the worldwide prevalence of human diphyllobothriasis was estimated at nine million cases.4 It is frequently reported from Japan, south Korea, the Baltic countries, Scandinavia, western, and eastern Russia, in North America (Pacific Northwest). Some reports suggested that this parasitosis may emerge or re-emerge in Europe,5 although South America and other countries are regions where sporadic cases had been reported so far.6 A recent review by Dick gives precise data on the geographical distribution of this parasitosis.2

56.1.1 Classification and Morphology Diphyllobothrium is a genus belonging to Family Diphyllobothriidae, Order Diphyllobothriidea7 (formerly Pseudophyllidea), Class Cestoda and Phylum Platyhelminthes. To date, many species have been described within the Diphyllobothrium genus but only a dozen have been reported as human pathogens; the most frequent species being D. latum, D. nihonkaiense, and D. dendriticum.1–3 However, some species are only quoted in review papers or book chapters but are not mentioned in bibliographic databases (Table 56.1). Species

which are regularly reported in humans are D. latum (cosmopolitan), D. dentriticum (northern part of the northern hemisphere), D. nihonkaiense (Japan, South Korea, eastern Russia and Canada) and D. pacificum (Peru, Chile). Infections with the species Diplogonoporus grandis have been reported in Japan and recent molecular studies8 revealed a close relationship with the cetacean tapeworm D. stemmacephalum. Other species (D. alascense, D. cameroni, D. cordatum, D. dalliae, D. hians, D. ursi, D. yonagoense, and D. lanceolatum) have been also described in humans but few details are available in the scientific literature.

56.1.2  Biology and Pathogenesis The life cycle of D. latum, probably the most frequent species in humans, is complex and involves several hosts.2,3 Released in water, the eggs mature within eight to 12 days at a water temperature of 16–20°C, and yield a coracidium larva that is ingested by a zooplanktonic copepod crustacean. About 40 copepod species of the Eudiaptomus or Cyclops genus are likely to be the first intermediate hosts. This larva develops into a procercoid larva within the general cavity of the copepod. When carnivore fish ingest planktonic crustaceans, the larva develops into a plerocercoid larva, a few millimeters long. It migrates into the fish musculature or viscera where it can remain inactive for several years, but can re-encyst several times in other predatory fish. In Europe, the types of fish susceptible to host the larvae are perch (Perca fluviatilis), pike (Esox lucius), charr (Salvelinus alpinus), and burbot (Lota lota). The Coregonidae and probably the Salmonidae of Salmo genus do not host D. latum larvae, but they can host D. dentriticum larvae. The Pacific Salmonidae of the genus Onchorynchus are the usual hosts for D. nihonkainese. 781

782

Molecular Detection of Foodborne Pathogens

Table 56.1 Species of Diphyllobothrium Reported from Humansa and Availability of Sequences in Genbank (January 2008) Species

18S

ITS

COI

NADH

D. alascense (Rausch et Williamson, 1958) D. cameronib (Rausch, 1969) D. cordatum (Cobbold, 1869)

Burbot, Smelt Marine fishes Marine fishes

Dogs Monk seal Seals, walrus

Alaska Japan North Atlantic

0 0 0

0 0

0 0 0

D. dalliae (Rausch, 1956) D. dendriticum* (Nitzsch, 1824)

Dallia pectoralis Salmonids, Coregonus

Gulls, dogs Fish eating birds

Alaska Circumpolar

0

 +  0

0 0 0 0

0

D. hiansb (Diesing, 1850) D. klebanovskiic (Muratov and Psokhov, 1988) D. lanceolatum (Krabbe, 1865)

Marine fishes Salmonids

Seals Brown bears

North Atlantic Eastern Eurasia,

 +  0

 +  0

 +  0

 +  0

 + 

 + 

 + 

 + 

Coregonus

Seals

North Pacific

0

 + 

0

0

D. latum* (L., 1758)

Pike, Burbots, Percids

Dogs, bears

Cosmopolitan

 + 

 + 

 + 

 + 

D. nihonkaiense*c (Yamane, 1986)

Salmon

?

North Pacific

 + 

 + 

 + 

D. pacificum* (Nybelin, 1931)

Marine fishes

Sea lions, fur seals

Peru, Chile

Red salmon

Bears

 +  0

D. yonagoense (Yamane, 1981) Diplogonoporus grandisd (Blanchard, 1889)

Salmon Anchovy, sardine

? Cetacean

Alaska, British Columbia Japan, Eastern Siberia Japan

 +  0

 +  0

D. ursi (Rausch, 1956)

 +  0

a

b c d

Fish Hosts

Other Hosts

Distribution

0

0

0

0

0

 + 

 + 

 + 

 + 

Source: Data summarized from Dick, T.A., In Foodborne Parasitic Zoonoses, Fish and Plantborne Parasites, Murrell, K.D. and Fried, B. (eds), Springer, New York, 2007; Chai, J.Y., Murrell, K.D. and Lymbery, A.J., Int. J. Parasitol., 35, 1233, 2005; Muller, R., In Worms and Human Disease, 2nd edn, CABI, Wallingford, UK, 2002. No mention of this species in PubMed bibliographic data base. D. nihonkaiense and klebanowskii could be synonymous according to sequences published by Arizono, N. et al., Parasitol. Int., 57, 212, 2008. This could be synonymous with D. stemmacephalum according to Arizono, N. et al., Parasitol. Int., 57, 212, 2008 and it is proved in humans by molecular studies.

D. pacificum and D. grandis larvae are hosted by marine fish. Man and other ichtyophagous animals become contaminated by ingesting raw fish. The plerocercoid larva can grow between 5 and 20 cm a day2 and develops into an adult that yields its first eggs about 1 month after infestation. D. latum is the longest human parasite known (about 10 m long) and can live for several years. Its symptoms, although limited, are polymorphous: manifestations may include abdominal discomfort (abdominal pain, diarrhea), weight loss, asthenia, and vertigo. Anaemia due to vitamin B-12 deficiency has been described in case of prolonged infestation.2,3 Human experimental infestations have been described.2 Three volunteers, infected by two to three plerocercoid larvae, did not present any obvious clinical symptoms except for the release of proglottis. The two non-\treated subjects dewormed spontaneously 7 months (in the first case), and 4 years and 6 months (in the second case) after being infected. The parasite is sensitive to praziquantel (15 mg/kg/day in one dose) and to niclosamide (2 g on an empty stomach in two doses an hour apart). In experimental infections, D. latum, D. dendriticum, D. ditremum, and D. vogeli demonstrate variations in their developmental patterns. Specifically, D. latum tends to shed the entire larval body easily with an average rate of 82.1% in both hamsters and rats; D. dendriticum displays a shedding rate of 8.7% in the hamsters and 34.9% in the rat; D. ditremum exhibits a shedding rate of 42.9%, but it is difficult to recover from hamsters; and D. vogeli has a recovery rate of 38.3% without shedding.34

56.1.3 Conventional Diagnostic Techniques All members of the genus Diphyllobothrium are morphologically very similar and are sometimes distinguished by hosts and geographical origin. The different species infective for humans can be distinguished by skillful parasitologists using morpho-anatomical criteria,9,10 but these criteria are relative as they vary with age, with the degree of development and physiological modifications.11 Diagnosis is possible on the whole worm, rarely obtained, showing a medium-sized to large tapeworm with strobila usually segmented. Genital apertures are on the ventral face and the scolex is unarmed with dorsal and longitudinal grooves, named bothria. On the opposite, proglottids are easily obtained from the feces of infected animals. They are wider than long with genital pores opening midventrally (Figure 56.1). Each segment measures 2–7 × 10–12 mm. Eggs are oval and operculated. Eggs of D. latum are greyish brown with a 72 μm average length and a 57 μm average width. Eggs of D. nihonkaiense are smaller (57 × 44 μm). However sizes vary considerably among individual worms and could decrease with intensity of infection.9 Eggs can be mistaken with eggs of Paragonimus sp or Fasciola hepatica whose sizes are larger. The plerocercoid larvae, obtained from fishes, are ranging from few millimeters to a few centimeters long and their “identification has always been the job of an expert”.10 They appear as white elongated masses in the skeletal muscle of

783

Diphyllobothrium

Figure 56.1  Proglottids of Diphyllobothrium latum. Note the ventral opening of the genital aperture.

Figure 56.2  Plerocercoid of Diphyllobothrium latum extracted from a perch filet.

certain fish families (Figure 56.2). They can be confounded by nonexperts with acanthocephales (easily distinguishable by the presence of an evertable proboscis armed with spines) or with cestodes of the genus Proteocephalus. Andersen and Gibson (1989) published keys to identify three species of larval Diphyllobothrium occurring in European and North American freshwater fishes.10 These keys are based on features of gross-morphology visible under a light or a scanning electron microscope, on body-length and site within the host and features visible in histological sections.

56.1.4 Molecular Diagnostic Techniques Biochemical techniques (isoenzymatic assay, or immuno-electrophoresis) have been used as alternatives to the traditional tools for species identification, but the development of molecular biology resulted in a better knowledge of the Diphyllobothrium genus.12–14 For example, Matsuura et al.14 utilized restriction fragment length polymorphisms (RFLPs) of ribosomal DNA (rDNA) to

discriminate between D. latum and D. nihonkaiense as the RFLP profiles generated with restriction enzymes SmaI, Hinf I, and HhaI provided valuable species-specific markers. Since 1998, interesting results in the study of the Diphyllobothrium genus have been obtained by sequences analysis of the 18S rRNA, cytochrome c oxidase subunit 1 (COI), NADH dehydrogenase subunit 3 genes and of the partial internal transcribed spacer 2 (ITS2) region.15–19 By using nucleotide sequence analysis of a region of the cytochrome c oxidase subunit I gene of mitochondrial DNA from D. nihonkaiense plerocercoids recovered, Ando et al.18 confirmed the first identification of D. nihonkaiense from Oncorhynchus masou ishikawae fish in Japan. Upon examination of sequences of the ITS-2 rRNA genes of the family Diphyllobothriidae, Logan et al.19 uncovered many phylogenetic insights among genera Ligula, Digramma, Diphyllobothrium, and Schistocephalus, including establishment of Schistocephalus as the most basal taxon of the Diphyllobothriidae, and ligulids (previously family Ligulidae) as being part of the Diphyllobothriidae.

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Molecular Detection of Foodborne Pathogens

Taenia saginata AF096240 Proteocephalus sp. SK RK3 RK4 AY972073 AY972071 BW1 BW2 FD AG ADB LF JN 02.05 JDC 04.04 JDC 02.04 JDC 11.03

D. stemmacephalum

D. ditremum

D. latum

AY972072 PCH CSS KCH AB015755 RK1

D. nihonkaiense

D. dendriticum

RK2 0.1

Figure 56.3  Phylogenetic construction based on partial COI gene sequences. Referent sequences obtained from databases: probable Taenia sp. specimen (AF096240); D. latum proglottids, French isolate (AY972073); D. latum larva, Leman lake Perca fluviatilis isolate (AY972071); Diphyllobothrium sp HY-2005 proglottids, French isolate (AY972072); D. nihonkaiense, Japanese isolate (AB015755). Significant bootstrap values were observed for each node. Details on the other isolates can be found in reference 20.

Recently, we determined the value of 18S rRNA gene, internal transcribed spacer 1 (ITS1) and COI gene sequences to differentiate between Diphyllobothrium isolates. Sequences from 18 isolates (larvae or adults) of D. latum, D. nihonkaiense, D. ditremum, D. dentriticum, and D. stemmacephalum species were examined. COI region sequence analysis was clearly more discriminative than those of the ITS1 and 18S rRNA and was a useful tool for identifying specimens.20 In our own experience, analysis of the 18S rRNA or ITS1 regions of different Diphyllobothrium species showed less divergence (2–6%) than the COI region (13–14%). The 18S rRNA gene has already been described as a very conserved genomic target, and analysis of its nucleotide sequence provides a wealth of information about phylogenetic relationships of biological organisms concerned.21 This is due to the fact that because 18S rRNA gene is present in multiple copies, contains regions of relative conservation as well as diversity and demonstrates patterns of concerted evolution that occurs among repeated copies. A recent study using ITS2 sequences showed that D. pacificum was clearly

different (with sequence similarity between 79.0 and 80.2%) from other members of the genus (e.g., D. cordatum, D. dendriticum, and D. lanceolatum) and that ITS2 could not discriminate between D. ditremum and D. dendriticum.22 Similarly, ITS1 sequences could not discriminate between D. ditremum and D. dendriticum.20 COI gene analysis provided the most discriminative sequences between the different species (Figure 56.3). The sequences of NADH dehydrogenase subunit 3 gene are promising to discriminate the different species, because sequences between D. latum, D. nihonkaiense, and D. dentriticum are more divergent than COI sequences.23 Mitochondrial sequences are known to be more polymorphic than the 18S rRNA and ITS1 nuclear sequences, as restriction endonuclease cleavage maps of the mitochondrial DNA from individual species reveal that the species differ from one another at most of the cleavage sites due partially to an elevated rate of mutation in mitochondrial DNA.24 Therefore, mitochondrial DNA can be also targeted for high-resolution analysis of the evolutionary process among Diphyllobothrium species.20

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Diphyllobothrium

56.2  Methods We provide here the technique to identify Diphyllobothrium samples (larval stages obtained from fish, eggs or proglottids obtained from patients or mammals) by amplification and DNA sequencing of mitochondrial targets (COI, NADH dehydrogenase) which are very useful to identify Diphyllobothrium species. However as these genes have not been sequenced for some species, nuclear targets such as 18S rRNA or ITS could still be useful.

56.2.1 Reagents and Equipments Reagents and equipment needed are listed in Table 56.2.

56.2.2 Sample Preparation Sample collection. Fish filets are examined for larvae by cutting the flesh in 2–3 mm thick slices. Proglottids are discharged with the feces. Eggs are concentrated from stools by usual diphasic methods. Larvae and proglottids are washed with 0.9% sodium chloride. Eggs are submitted to three cycles of washing with 0.9% sodium chloride and centrifugation 5 min at 200 × g (300 rpm). One larva, one proglottid, or the pellet of feces concentration is used for analysis. All samples must be stored in 100% ethanol at –20°C before analysis and DNA extraction. DNA extraction from samples. Proglottids can be pretreated by three cycles of freezing at −80°C and defreezing at 60°C before DNA extraction,23 but this pre-treatment is not compulsory.20 DNA from the different samples is extracted by using QIAamp DNA Mini kit with the tissue protocol (Qiagen, France) according to the manufacturer's instruction. Twenty microliters of proteinase K are mixed with the sample and 180 µl of buffer ATL and incubated for 1 h (larvae) or 4 h (proglottids, eggs) at 56°C. Two hundred microliters of buffer AL are then added and the sample is incubated for 10 min at 70°C. Two hundred microliters of 100% ethanol are added and DNA extracted with the use of the QIAamp column and centrifugation for 1 min at 6000 × g. DNA is washed with 500 µl of buffer AW1 (eliminated by a 1 min

centrifugation at 6000 × g) then with 500 µl of buffer AW2 (eliminated by a 5 min centrifugation at 17900 × g). A last centrifugation is performed for 1 min at 17900 × g to eliminate residual buffer. DNA is then eluted by the addition of 100 µl sterile distilled water in the column, a 5 min incubation and centrifugation for 1 min at 6000 × g.

56.2.3  Detection Procedure Principle: The partial region of the nuclear 18S rRNA gene is amplified using primers 81 and 83, designed for Eucestoda molecular phylogenetic study.15 The nuclear internal transcribed spacer regions (ITS1 and ITS2) including 5.8S rRNA gene are amplified using primers BD1 and BD2, originally designed for flukes analysis.25 The mitochondrial COI gene is amplified by using JB3 and JB4.5 primers designed to analyse Echinoccocus sp.26 and previously used to identify Diphyllobothrium species.23 As for some specimens, no amplification was obtained, we designed new primers: JB6 and JB5R targeting a larger region. The mitochondrial tRNA-Pro, tRNA-Ile, tRNA-Lys, NADH dehydrogenase subunit 3 genes are amplified using the primers DnND3Fo and DnND3Re, designed to identify Diphyllobothrium species.23 All the primers are described in Table 56.3. Procedure: (1) Prepare PCR mixture (50 µl) by adding 10 µl of DNA, 5 µl of High Fidelity Platinium DNA polymerase 10 × buffer (Invitrogen, France), 1.5 or 4 µl of 50 mM of MgSO4, 10 nmol of each DNTP (dATP, dTTP, dCTP, dGTP), 10 or 40 pmol of each primer and 1 or 3 U of High Fidelity Platinium DNA polymerase (Invitrogen), and sterile distilled water (to a final volume of 50 µl). A tube containing 10 µl of sterile water in place of 10 µl of DNA is included as a negative control. (2) Apply the following cycling program in a thermal cycler: a denaturation phase at 94°C for 7 min, different cycles of amplification according to the target (see Table 56.4) and a final elongation cycle of 10 min at 68°C.

Table 56.2 Reagents and Equipment Stage

Reagents

DNA extraction

QIAmp DNA Mini kit (Qiagen, France) thanol 100%

PCR

High Fidelity Platinium DNA polymerase (Invitrogen, France) Each DNTP 25 µmoles (GE Healthcare, France) Seaken GTC Agarose (Tebu, France) Tris/Acetate/EDTA (TAE) buffer (Gibco, France) Ethidium bromure (Eurobio, France) Molecular weight VI (Roche Diagnostics) QIAquick PCR Purification kit (Qiagen)

Genotyping



Supplies and Equipment Laminar flow cabinet Dry bath incubator Refrigerated microtube centrifuge Vortex agitator Microtube centrifuge Thermal Cycler (Applied Biosystem, France) Gel electrophoresis apparatus Ultraviolet lamp Refrigerated microtube centrifuge Automated DNA sequencer

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Molecular Detection of Foodborne Pathogens

Table 56.3 Primers Used for Diphyllobothrium Species Identification Target

Type

Primers

18S rRNA

Nuclear

83: 5′-GATACCGTCCTAGTTCTGACCA 81: 5′-TTCACCTACGGAAACCTTGTTACG

ITS1 & ITS2

Nuclear

BD1: 5′-GTCGTAACAAGGTTTCCGTA BD2: 5′-TATGCTTAARTTCAGCGGGT

Cytochrome c oxidase subunit 1

Mitochondrial

JB3: 5′-TTTTTTGGGCATCCTGAGGTTTAT JB4.5: 5′- TAAAGAAAGAACATAATGAAAATG JB6: 5′-GATAGTAAGGGTGTTGA JB5R: 5′-CAAGTATCRTGCAAAATATTATCAAG

NADH dehydrogenase subunit 3

Mitochondrial

DnND3Fo: 5′-CGTTAAGATGAATGAG DnND3Re: 5′-GACCTTACATCACTTAGTG

Table 56.4 Magnesium and Primers Concentrations, DNA Polymerase Units, Amplification Cycles and Length of Amplicons for Diphyllobothrium Species Identification 50 mM MgSO4

Primer Concentration

DNA Polymerase

83 and 81

1.5 µl

10 pmol

1U

BD1 and BD2

1.5 µl

10 pmol

1U

JB3 and JB4.5

4 µl

40 pmol

3U

JB6 and JB5R

1.5 µl

10 pmol

1U

DnND3Fo and DnND3Re

4 µl

40 pmol

3U

Primers

(3) Analyze 15 µl of amplicons in 1 or 2% agarose gels containing 0.5 µg/ml ethidium bromide and visualize bands with a UV illuminator. (4) Purify the remaining 35 µl amplicons using a PCR purification kit (e.g., QIAquick PCR Purification kit, Qiagen) and sequence in both directions using an automated DNA sequencer. Namely, mix 35 µl of positive PCR amplicons with five volumes of buffer PBI; extract DNA with QIAquick column; centrifuge for 1 min at 17900 × g; wash DNA with 750 µl of buffer PE, which is eliminated by two centrifugations of 1 min at 17900 × g; elute DNA by adding 50 µl sterile distilled water in the column and after a 5 min incubation and centrifugation for 1 min at 17900 × g. (5) Mix 10 ng per 100 bp of each purified amplimer with 4 picomol of the corresponding forward or reverse primers and analyze in an automatical sequencer. (6) Align resulting sequences with Clustal W, make minor corrections by using ChromasPro 1.32 (Technelysium) and Bioedit 5.0, and BLAST the Diphyllobothrium sequences into DDBJ/EMBL/ GenBank databases where sequences are available for most species (see Table 56.1).

Cycling Program 38 cycles of 94°C/30 sec, 55°C/40 sec, and 68°C/1 min 30 sec 41 cycles of 94°C/30 sec, 50°C/40 sec and 68°C/40 sec 41 cycles of 94°C/30 sec, 50°C/40 sec, and 68°C/1 min 38 cycles of 94°C/30 sec, 55°C/40 sec, and 68°C/40 sec

Amplicon (bp) 1000 1100 440 650 708

56.3 Conclusions and Future Perspectives The methods described above have been very useful for the typing of parasitic specimens obtained from fish,18,27 or human stools,18,20,23,28,29 and will certainly allow a more precise classification of the genus. The growing popularity of raw fish dish has probably led to an increase of diphyllobothriasis in Europe and elsewhere,5,6 Imported Pacific salmon or travel in endemic countries have led to the occurrence of cases of D. nihonkaiense or D. dendriticum in France and Switzerland,23,28,29 Therefore, it is of importance, owing to the high morphologic similarities between these species, to precisely identify all isolates. In our own experience, amplification of the partial COI gene was obtained with the JB3-JB4.5 primers from all samples except for D. stemmacephalum. For the latter species, DNA amplification is possible when the JB5-JB6 primers are used. In addition, recent studies by Arizono et al. showed that this species could be synonymous with Diplogonoporus grandis.26 D. dendriticum and D. ditremum could be differentiated only by using the mitochondrial sequence.20 Both species are sharing the same geographic areas (circumpolar), the same intermediate (coregonids) and definitive hosts (fish eating birds and mammals).

787

Diphyllobothrium

However, only D. dendriticum is a human pathogen. This approach is also useful for confirming the COI differences between D. nihonkaiense and D. latum.17,23 The COI gene is the most appropriate target to identify specimens and to classify species described in humans, because DNA sequences are enough divergent between species and are available in databases for most species. Moreover, the recent determination of the mitochondrial genomes of D. latum and D. nihonkaiense may provide additional tools to study intraspecific variations.30 The sequences of NADH dehydrogenase subunit 3 gene are promising to discriminate the different species, because sequences between D. latum, D. nihonkaiense and D. dentriticum are more divergent than COI sequences (13% vs 8%). Mitochondrial sequences are known to be more polymorphic than the 18S rRNA and ITS1 nuclear sequences. In 2006, we identified the first putative D. nihonkaiense infection in France from the consumption of an imported Pacific salmon from Canada.23 D. nihonkaiense was first described in 1986, when Yamane et al. showed morphological and biochemical differences between Finnish and Japanese isolates of D. latum; the latter was renamed D. nihonkaiense.18 This parasite is now recognized as a frequent cause of diphyllobothriasis in Japan, contracted by the consumption of raw Pacific salmon (Oncorhynchus keta or O. masou). Along the Pacific coast of Russia (the Khabarovsk Territory, the Sakhalin and Kamchatka regions), populations are frequently infected with D. klebanovskii, which is found in O. keta and O. gorbuscha muscle tissue.31 Diphyllobothriasis has also been described among Oncorhynchus consumers along the western coast of the United States,32 and a significant upsurge was observed in the late seventies (17 cases in 1979, 64 in 1980). D. ursi, usually infecting bears, has been reported among Alaskan residents following Oncorhynchus consumption.33 The migratory routes of salmon of Asian and American origin overlap in the north Pacific and we suggested in 2006 that the reported absence of D. nihonkaiense along the Pacific coast of North America might be in fact due to its misidentification. A second D. nihonkaiense infestation was also reported in Switzerland in 2007.29 Very recently, sequences published by Arizono et al. (accession numbers AB375660-AB375662, AB375672, and AB375673) showed that D. klebanowkii from eastern Russia was similar to the Korean and Japanese D. nihonkaiense. Typing D. ursi isolates would be now, very interesting to be sure that this species is not D. nihonkaiense. This example illustrates the fact that taxonomy in the genus Diphyllobothrium will evolve now very rapidly with the new tools provide by molecular biology. Sequencing is a certainly precise approach to type specimens of Diphyllobothrium, but it is only feasible with species whose related gene sequences are in the databases.



















References

1. Chai, J.Y., Murrell, K.D. and Lymbery, A.J. Fish-borne parasitic zoonoses: status and issues. Int. J. Parasitol., 35, 1233, 2005.



2. Dick, T.A. Diphyllobothriasis: the Diphyllobothrium latum human infection conundrum and reconciliation with a worldwide zoonosis. In Foodborne Parasitic Zoonoses, Fish and Plantborne Parasites, pp. 151–184. Murrell, K.D. and Fried, B. (eds). Springer, New York, 2007. 3. Muller, R. The cestodes. In Worms and Human Disease, 2nd edn, pp. 63–105. CABI, Wallingford, UK, 2002. 4. Crompton, D.W.T. How much human helminthiasis is there in the world. J. Parasitol., 85, 397, 1999. 5. Dupouy-Camet, J. and Peduzzi, R. Current situation of human diphyllobothriasis in Europe. Euro Surveill., 9, 5, 2004. 6. Sampaio, J.L. et al. Diphyllobothriasis, Brazil. Emerg. Infect. Dis., 11, 1598, 2005. 7. Kuchta, R. et al. Suppression of the tapeworm order Pseudophyllidea (Platyhelminthes: Eucestoda) and the proposal of two new orders, Bothriocephalidea and Diphyllobothriidea. Int. J. Parasitol., 38, 49, 2008. 8. Arizono, N. et al. Diplogonoporiasis in Japan: genetic analyses of five clinical isolates. Parasitol. Int., 57, 212, 2008. 9. Andersen, K. and Halvorsen, O. Egg size and form as taxonomic criteria in Diphyllobothrium (Cestoda, Pseudophyllidea). Parasitology, 76, 229, 1978. 10. Andersen, K. and Gibson, D. A key to three species of larval Diphyllobothrium Cobbold, 1858 (Cestoda: Pseudophyllidea) occurring in European and North American freshwater fishes. Syst. Parasitol.,13, 3, 1989. 11. Dick, T.A. and Poole, B.C. Identification of Diphyllobothrium dendriticum and Diphyllobothrium latum from freshwater fishes of central Canada. Can. J. Zool., 63, 196, 1985. 12. Fukumoto, S. et al. Distinction between Diphyllobothrium nihonkaiense and Diphyllobothrium latum by immunoelectrophoresis. Jpn J. Parasitol., 37, 91, 1988. 13. De Vos, T. and Dick, T.A. Differentiation between Diphyllobothrium dendriticum and Diphyllobothrium latum using isoenzymes, restriction profiles and ribosomal gene probes. Syst. Parasitol., 13, 161, 1989 14. Matsuura, T., Bylund, G. and Sugane, K. Comparison of restriction fragment length polymorphisms of ribosomal DNA between Diphyllobothrium nihonkaiense and D. latum. J. Helminthol., 66, 261, 1992. 15. Mariaux, J. A molecular phylogeny of the eucestoda. J. Parasitol., 84, 114, 1998. 16. Olson, P.D. and Caira, J.N. Evolution of the major lineages of tapeworms (Platyhelminthes: Cestoda) inferred from 18S ribosomal DNA and elongation factor-1. J. Parasitol., 85, 1134, 1999. 17. Isobe, A. et al. Molecular phylogeny of Diphyllobothrium nihonkaiense (Yamane et al., 1986) and other diphyllobothiid tapeworms based on mitochondrial cytochrome c oxidase subunit I gene sequence. Parasitol. Int., 47 (suppl 1), 138, 1998. 18. Ando, K. et al. Five cases of Diphyllobothrium nihonkaiense infection with discovery of plerocercoids from an infective source, Oncorhynchus masou ishikawae. J. Parasitol., 87, 96, 2001. 19. Logan, F.J. et al. The phylogeny of diphyllobothriid tapeworms (Cestoda: Pseudophyllidea) based on ITS-2 rDNA sequences. Parasitol. Res., 94, 10, 2004. 20. Yera, H., Nicoulaud, J. and Dupouy-Camet, J. Use of nuclear and mitochondrial DNA PCR and sequencing for molecular identification of Diphyllobothrium isolates potentially infective for humans. Parasite, 15, 402, 2008. 21. Hillis, D. and Dixon, M. Ribosomal DNA: Molecular evolution and phylogenetic inference. Quart. Rev. Biol., 66, 411, 1991.

788 22. Skerikova, A. et al. Is the human-infecting Diphyllobothrium pacificum a valid species or just a South American population of the holarctic fish broad tapeworm, D. latum? Am. J. Trop. Med. Hyg., 75, 307, 2006. 23. Yera, H. et al. Putative Diphyllobothrium nihonkaiense acquired from a Pacific salmon (Oncorhynchus keta) eaten in France; genomic identification and case report. Parasitol. Int., 55, 45, 2006. 24. Brown, W.M., George, M. Jr. and Wilson, A.C. Rapid evolution of animal mitochondria DNA. Proc. Natl. Acad. Sci. USA, 76, 1967, 1979 25. Luton, K., Walker, D. and Blair D. Comparisons of ribosomal internal transcribed spacers from two congeneric species of flukes (Platyhelminthes: Trematoda: Digenea). Mol. Biochem. Parasitol., 56, 323, 1992. 26. Bowles, J., Blair, D. and McManus D.P. Genetic variants within the genus Echinococcus identified by mitochondrial DNA sequencing. Mol. Biochem. Parasitol., 54, 165, 1992. 27. Nicoulaud, J., Yera, H. and Dupouy-Camet, J. Prevalence of Diphyllobothrium latum, L., 1758 infestation in Perca fluviatilis from Lake Leman. Parasite, 12, 362, 2005 (in French).

Molecular Detection of Foodborne Pathogens 28. Wicht, B., de Marval, F. and Peduzzi, R. Diphyllobothrium nihonkaiense (Yamane et al., 1986) in Switzerland: first molecular evidence and case reports. Parasitol. Int., 56, 195, 2007. 29. Wicht, B. et al. Imported diphyllobothriasis in Switzerland: molecular evidence of Diphyllobothrium dendriticum (Nitsch, 1824). Parasitol. Res., 102, 201, 2008. 30. Nakao, M. et al. Mitochondrial genomes of the human broad tapeworms Diphyllobothrium latum and D. nihonkaiense (Cestoda: Diphyllobothriidae). Parasitol. Res., 101, 233, 2007. 31. Muratov, I.V. and Posokhov, P.S. Causative agent of human diphyllobothriasis Diphyllobothrium klebanovskii sp. Nov. Parazitologia, 22, 165, 1988. 32. Ruttenber, A.J. et al. Diphyllobothriasis associated with salmon consumption in Pacific Coast states. Am. J. Trop. Med. Hyg., 33, 455, 1984. 33. Rausch, R.L. and Hilliard D.K. Studies on the helminth fauna of Alaska. XLIX. The occurrence of Diphyllobothrium latum (Linnaeus, 1758) (Cestoda: Diphyllobothriidae) in Alaska, with notes on other species. Can. J. Zool., 48, 1201, 1970. 34. Yamane, Y. et al. Early development of four Diphyllobothrium species in the final host. Parasitol. Res., 74, 463, 1988.

57 Fasciola

Xing-Quan Zhu, Qing-Jun Zhuang, and Rui-Qing Lin South China Agricultural University

Wei-Yi Huang

Guangxi University

Contents 57.1 Introduction.................................................................................................................................................................... 789 57.1.1 Classification and Morphology of Fasciola spp. ............................................................................................. 789 57.1.2 Biology, Pathogenesis, and Medical Importance . ........................................................................................... 789 57.1.3 Laboratory Identification and Diagnosis ......................................................................................................... 790 57.1.3.1 Differentiation of Fasciola spp. from Definitive Hosts................................................................... 790 57.1.3.2 Detection of Fasciola Infection in Snails by PCR.......................................................................... 790 57.2 Methods.......................................................................................................................................................................... 791 57.2.1 Reagents and Equipment................................................................................................................................... 791 57.2.2 Sample Preparation and DNA Isolation............................................................................................................ 791 57.2.3 Detection Procedures........................................................................................................................................ 791 57.3 Conclusion and Future Perspectives............................................................................................................................... 792 References.................................................................................................................................................................................. 792

57.1  Introduction 57.1.1 Classification and Morphology of Fasciola spp. The causative agents of human and animal fascioliasis, Fasciola spp., belong to the family Fasciolidae, subclass Digenea, class Trematoda. Several species have been described within the genus Fasciola, but only two species, Fasciola hepatica and Fasciola gigantica, are commonly recognized as taxonomically valid occurring in animals and humans,1,2 with F. hepatica mainly occurring in temperate areas, F. gigantica in tropical zones, and both overlapping in subtropical areas.2,3 Adults of both species occur in many domestic ruminants and humans and can cause serious disease.2,4 The two species differ in pathological and epidemiological characteristics.2 In addition to F. hepatica and F. gigantica, several recent studies using the first and/or second internal transcribed spacers (ITS-1 and ITS-2) of ribosomal DNA (rDNA) as genetic markers have identified an “intermediate form” of Fasciola between F. hepatica and F. gigantica from Japan, Korea, China, and Vietnam.5–12 Fasciola spp. are large leaf-shaped flukes, this morphological feature and their predilection site in hosts can be used for their tentative identification. The young F. hepatica at the time of entry into the liver is 1.0–2.0 mm in length

and lancet-like. When it has become fully matured in the bile ducts, it is leaf-shaped, grey-brown in color and measures around 3.5 cm in length and 1.0 cm in width. The anterior end is conical and marked off by distinct shoulders from the body. The tegument is covered with backwardly projecting spines. An oral and ventral sucker may be readily seen. The egg is oval, operculate, yellow, and large (around 150 × 90 µm), and approximately twice the size of a trichostrongyle egg.13 F. gigantica is larger than F. hepatica in size and can reach 7.5 cm in length. Its shape is more leaf-like, but the concial anterior end is very short and the shoulders characteristic of F. hepatica are barely perceptible. The eggs are larger than those of F. hepatica, measuring 190 × 100 µm.13

57.1.2 Biology, Pathogenesis, and Medical Importance Biology. The life cycle of F. hepatica consists of seven stages, namely egg, miracidium, sporocyst, redia, cercaria, metacercaria, and adult fluke. The development of the parasite requires two hosts (intermediate host and definitive host).2,13 Mammals are usually definitive hosts, infected through ingesting infective metacercariae, with sheep and cattle being the most important. The metacercariae excyst in the small intestine, penetrate the intestinal wall, move into

789

790

the peritoneal cavity, and invade into the liver parenchyma through the capsule. About 6 weeks later, the parasites finally settle down in the biliary tract where they become mature and produce eggs that are excreted with stools. The eggs can hatch into the ciliated miracidia that can invade their specific snail intermediate hosts to undergo asexual reproduction.13 After several developmental stages, the ciliated miracidia can gradually evolve into metacercariae with infectivity on the leaves of water plants or the surface of other organisms. A new life cycle then begins. The life cycle of F. gigantica is similar to that of F. hepatica, the main differences being in the time scale of the cycle. Most parasitic phases are longer and the prepatent period is 13–16 weeks.13 The intermediate hosts of Fasciola spp. are freshwater snails of the genus Lymnaea. There are more than 20 species in the genus, such as L. columella, L. viatrix and L. cubensis, many of which transmit F. hepatica and F. gigantica.13 Pathogenesis. Fasciola spp. are highly pathogenic. In humans, pathogenesis depends on the number of infecting flukes, and appears to be similar to that reported in animals.14–17 F. hepatica has a special propensity for the liver and can eventually migrate into the biliary ducts. The fascioliasis disease process often shows two phases: acute phase and chronic phase. In the acute phase, the liver is invaded by the immature flukes, which cause the patient to have a fever, hepatomegaly, abdominal pain, eosinophilia, urticaria, hemobilia, and hepatica nodules.18,19 In the chronic phase, the parasite exists in the biliary ducts and usually obstructs them, which can cause jaundice, wasting, anemia, ascites, cardiac disorder, and pancreatitis.18–20 However, a variety of symptoms and signs can arise in other organs or systems apart from the liver. These clinical manifestations may be caused by the larvae that accidentally lose their way to the liver and so reside in other organs or systems, which results in ectopic fascioliasis.19 The common ectopic locations of fascioliasis include the subcutaneous tissues and the lymph nodes,19,21 and the less frequent ectopic locations are the lung, eye, brain, stomach, cecum, epididymis, and peritoneum.22–24 The pathogenesis for these disorders is poorly understood, but the most possible mechanism is an immunoallergic interplay.19 Medical importance. Fascioliasis is a major parasitic disease of livestock with over 700 million production animals being at risk of infection and worldwide economic losses estimated at > US$3.2 billion per year.25,26 Human infection has been reported worldwide,27 and approximately 2.4 million people in more than 60 countries are estimated to be infected through ingesting freshwater plants or water containing metacercariae of Fasciola spp.28 and the number of people at risk is more than 180 million globally.16,29,30 In fact, fascioliasis has been considered an important plant- and water-borne parasitic zoonosis.31 Infected animals are the main source for human infection.

Molecular Detection of Foodborne Pathogens

57.1.3 Laboratory Identification and Diagnosis 57.1.3.1 Differentiation of Fasciola spp. from Definitive Hosts Direct parasitological techniques, indirect immunological tests and other noninvasive diagnostic techniques have been utilized for diagnosis of human fascioliasis. Coprological examination of eggs has been the traditional approach for the diagnosis of human and animal infection with Fasciola spp.13 Various immunological tests have been used for the diagnosis of human fascioliasis because they provide the advantages of being applicable during all stages of the disease, especially during the invasive or acute phase, and in other situations in which coprological techniques are inappropriate. Almost all of these techniques concern the detection of circulating antibodies and only a few are designed to detect circulating antigens and immune complexes,32 as well as stool antigen.33 Serological techniques have also recently proved to be useful for monitoring post-treatment evolution.34 However, immunological techniques may suffer from poor sensitivity and specificity.35 Noninvasive diagnostic techniques, which can be used for human diagnosis, include radiology, ultrasound, computer tomography and magnetic resonance.36,37 Molecular approaches have been used for the identification and differentiation of the causative agents of human and animal fascioliasis, because the differential diagnosis between F. hepatica and F. gigantica in humans cannot be easily achieved by clinical, pathological, coprological, or immunological methods. This is particularly problematic in those areas of overlapping range since differential diagnosis is very important due to the different pathological, transmission, and epidemiological characteristics of the two fasciolids. To distinguish between F. hepatica and F. gigantica, a simple and rapid PCR linked restriction fragment length polymorphism (PCR-RFLP) assay, using the common restriction enzymes Ava II and Dra II, has been described.38 It is based on a 618-bp long sequence of the 28S rDNA, provides unambiguous results and may be useful for both individual subject diagnosis and epidemiological surveys of humans and animals in endemic regions of sympatry in Africa and Asia.38 A similar PCR-RFLP assay using restriction endonucleases Hsp92 II and Rca I has recently been applied to differentiate between F. hepatica, F. gigantica and the “intermediate” Fasciola in China utilizing the second internal transcribed spacer (ITS-2) of rDNA.7 Using the first internal transcribed spacer (ITS-1) of rDNA as genetic marker, a recent study established a PCR-linked single-strand conformation polymorphism (PCR-SSCP) approach for the accurate identification and differentiation between F. hepatica, F. gigantica and the “intermediate form” of Fasciola in China.11 57.1.3.2 Detection of Fasciola Infection in Snails by PCR As the conventional techniques used for the detection of intra-molluscan stages of Fasciola species suffer from low

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Fasciola

sensitivity and specificity, the development of a highly sensitive and specific method for detecting infected snails would greatly aid in the epizootiological studies on F. gigantica or F. hepatica, by enabling the generation of accurate data on infection rates in field populations of snails. Recently, specific PCR assays were developed to detect Fasciola sp. infections in the snail intermediate hosts, utilizing various genetic markers, such as cytochrome c oxidase subunit 1 gene (cox1), ITS, and noncoding repetitive DNA fragment. Using forward primer 5′-ATTCACCCATTTCTGTTAGTCC-3′ and reverse primer 5′-ACTAGGCTTAAACGGCGTCC-3′) in standard PCR followed by sequence alignment, Krämer, and Schnieder39 demonstrated the sequence heterogeneity within a 120-bp repetitive DNA fragment in F. hepatica and F. gigantica. Similarly, Mostafa et al.40 also employed this primer pair for sensitive and specific detection of F. gigantica in Lymnaea natalensis. In addition, Velusamy et al.41 also utilized this primer pair for identification of F. gigantica infection in the snail intermediate host Lymnaea auricularia. The specific primers amplified a F. gigantica specific 124 bp noncoding repetitive DNA fragment from infected L. auricularia snails. Caron et al.42 examined Galba ­truncatula, Radix labiata and Radix balthica experimentally infected with F. hepatica by PCR and showed that R. labiata can act as an incidental intermediate host for F. hepatica. Cucher et al. developed a PCR assay targeting the cytochrome c oxidase subunit 1 gene.43 Using forward primer 5′-TATGTTTTGATTTTACCCGGG-3′ and reverse primer 5′-ATGAGCAACCACAAACCATGT-3′, these authors successfully amplified a 405-bp fragment from F. hepatica infected Lymnaea columella and Lymnaea viatrix. Magalhaes et al.44 designed a multiplex PCR assay for detection of F. hepatica DNA in L. viatrix snails which were formalin-fixed and paraffin-embedded. Specifically, the assay included a pair of primers targeting mitochondrial DNA (mDNA) for recognition of F. hepatica and another pair targeting the ITS region of the trematodes and the mollusc rDNA, to function as as an internal control. More recently, real-time PCR has been also developed for identification of Fasciola from intermediate snail host. Schweizer et al.45 utilized TaqMan chemistry that is based on the 5′–3′ exonuclease activity of the Taq DNA polymerase to cleave fluorescent dye-labeled TaqMan probe during PCR. The intensity of fluorescence that is proportional to the amount of PCR product formed is then measured by a sequence detection system. A combined use of primers and probe targeting an 86-bp long target of a repetitive 449-bp long genomic DNA fragment facilitated detection of Lymnaea truncatula naturally infected with F. hepatica. Overall, the PCR techniques are highly specific and sensitive and possess fairly good prospects of their utility as epidemiological tools for ascertaining the infectivity status in ubiquitous snail populations as well as for assessing host suitability for the parasites. But unfortunately, so far there had been no report of specific PCR assays which could allow the identification and differentiation of F. hepatica, F. gigantica and the “intermediate Fasciola.”

57.2  Methods 57.2.1 Reagents and Equipment Reagents Taq DNA polymerase (Takara) Agarose: (Sigma) Protease K (Merk) WizardTM DNA Clean-Up System Kit (Promega) Wizard™ PCR-Preps DNA Purification System Kit (Promega) Ethidium bromide (Promega) DNA size markers (Takara) Restriction enzyme Hsp92II and RcaI (Promega or Takara).



Equipment

Thermocycler (Whatman Biometra) Biophotometer (Eppendorf) Gel Documentation System (UVITEC) Bio-Rad Agarose Gel Electrophoresis System (­Bio-Rad) Desk centrifuge Pippets (2 µl/20 µl/200 µl/1000 µl) (Gilson)

57.2.2 Sample Preparation and DNA Isolation Adult trematodes of Fasciola can be collected from the liver of infected animal hosts at necropsy. Individual flukes were washed extensively in physiological saline, identified morphologically to genus and/or species according to existing keys and descriptions,1 and then fixed in 70% ethanol until extraction of genomic DNA. Total genomic DNA can be extracted from a portion of individual specimens by a method of sodium dodecyl sulphate/proteinase K (Merck) treatment, then columnpurified using WizardTM DNA Clean-Up System (Promega) and eluted into 60 µl H2O by following the manufactur­ er’s recommendations. DNA was also isolated from liver from cattle, buffalo, or goat using the same method as for ­t rematode samples. DNA samples were stored at -20°C until use. Fasciola DNA in formalin-fixed and paraffin-embedded Lymnaea tissues can be extracted by a technique reported by Magalhães et al.44 This technique includes deparaffinizing the slides with xylol, enzymatic digestion with proteinase K, purification with phenol chloroform and precipitation with isopropanol.

57.2.3  Detection Procedures Principle: Previous reports have demonstrated the value of targeting ITS-1 and ITS-2 of nuclear rDNA for molecular identification and phylogenetic investigations of ­platyhelminthes such as trematodes.9,46–48 The ITS-2 rDNA has proven especially useful for the genetic discrimination

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between F. hepatica and F. gigantica5,6,9,47,49 and for the confirmation of an intermediate genotype that differs from F. hepatica and F. gigantica.6 In light of these findings, Huang et al.7 designed a PCR-RFLP assay on the basis of genetic differences in the ITS-2 sequence for characterization and differentiation of Fasciola spp. In this assay, the ITS-2 rDNA from Fasciola spp. is amplified by PCR with forward and reverse primers flanking 5.8S and 28S sequences (i.e., forward primer 3S: 5′-GGTGGATCACTGGGCTCGTG-3′ and reverse primer BD2: 5′-TATGCTTAAATTCAGCGGGT-3′). The resulting PCR products are then used for the enzymatic digestion for differentiation of F. hepatica, F. gigantica and the “intermediate form” of Fasciola by using restriction enzyme Hsp92 II or Rca I. Procedure: (1) Prepare PCR mixture (100 µl) containing 1 × PCR buffer (10 mM Tris-HCl pH 8.4, 50 mM KCl, 2 mM MgCl2), 200 µM each of dNTP, 50 pmol of each primer (3S and BD2, see above), 2 U Taq polymerase (Promega), 1 µl purified Fasciola genomic DNA (approximately 10–20 ng) and distilled water (to 100 µl). Include tubes containing PCR mixture with host DNA or without genomic DNA in each PCR run as ‘negative’ controls. (2) Conduct PCR amplification in a thermocycler (Biometra or other brand) under the following conditions: one cycle of 95°C for 5 min; 30 cycles of 95°C for 1 min, 55°C for 1 min, and 72°C for 1 min; one cycle of 72°C for 10 min. (3) Examine an aliquot (5 µl) of each amplicon on 1.2% agarose-TBE (65 mM Tris-HCl, 22.5 mM boric acid and 1.25 mM EDTA, pH 9.0) gels. Include a DNA size marker (Takara) in a spare lane for estimation of the sizes of the ITS-2 amplicons. (4) Stain the gel with ethidium bromide (0.5 µg/ml) and photograph upon transillumination using a gel documentation system (UVITEC). (5) Purify the ITS-2 PCR products using spin columns (Wizard™ PCR-Preps DNA Purification System, Promega) following the manufacturer’s protocols. (6) Digest the purified Fasciola ITS-2 PCR products (5–15 µl, depending on DNA concentration) directly with 10 U (1 µl) of restriction enzyme Hsp92 II or Rca I in 20 µl for 4 h or overnight at 37°C, following the manufacturer’s recommendations. (7) Separate restriction fragments on 2.5% agarose gels. Also include a DNA size marker (Takara) in a spare lane. Stain and photograph the gel as in step 4. Note: On agarose gel, a fragment of approximately 550 bp in length will be amplified from each Fasciola genomic DNA and in no case is product amplified from no-DNA or host DNA sample. When the purified ITS-2 PCR products are digested with Hsp92 II, three bands of approximately 90 bp, 110 bp and 230 bp are produced for F. hepatica samples, whereas four bands of approximately 70 bp, 90 bp, 100 bp,

Molecular Detection of Foodborne Pathogens

and 160 bp are produced for F. gigantica samples, and five bands of approximately 70 bp, 90 bp, 100 bp, 160 bp, and 230 bp are produced for samples representing the “intermediate form” of Fasciola, allowing the identification and differentiation of F. hepatica, F. gigantica and the “intermediate form” of Fasciola.7 When the restriction endonuclease Rca I is used, the purified ITS-2 PCR products of F. hepatica samples remained undigested because they lack restriction site for Rca I, whereas F. gigantica samples will produce two bands of approximately 170 bp and 380 bp, respectively; and samples representing the “intermediate form” of Fasciola will display three bands of approximately 170 bp, 380 bp, and 550 bp, respectively, allowing the identification and differentiation of F. hepatica, F. gigantica and the “intermediate form” of Fasciola.7 This PCR-RFLP assay has been employed for the unequivocal delineation of the Fasciola spp. from Mainland China, and offers a valuable tool for the molecular identification and for studying the ecology and population genetic structures of Fasciola spp.

57.3 Conclusion and Future Perspectives Fascioliasis caused by the liver flukes of the genus Fasciola is a significant foodborne parasitic zoonosis in a number of countries, and millions of people are estimated to be infected throughout the world.2,16,25,29–31 The accurate identification and differentiation of Fasciola spp., and the diagnosis of human infection with Fasciola spp. is the prerequisite for the effective prevention and control of human fascioliasis. PCR-based approaches, such as PCR-RFLP, PCR-SSCP, and specific PCR assays, have been developed in the last decades for the accurate identification of Fasciola spp.7,11,39–45 These methods can also aid in the epizootiological studies of Fasciola spp., by enabling the generation of accurate data on infection rates in field populations of snails. Given that F. gigantica and F. hepatica can occur concurrently (overlapping) in subtropical areas, and the existence of the “intermediate form” of Fasciola in China, Japan, Korea, and Vietnam, there is a need to develop specific PCR assays for the identification and differentiation of F. gigantica, F. hepatica and the “intermediate form” of Fasciola in the near future.

References

1. Yamaguti, S. Systema helminthum. In The Digenetic Trematodes of Vertebrates, Vol. I. Interscience Publishers, Inc., New York, 1958. 2. Mas-Coma, S., Bargues, M.D., and Valero, M.A. Fascioliasis and other plant-borne trematode zoonoses. Int. J. Parasitol., 35, 1255, 2005. 3. Krämer, F., and Schnieder, T. Sequence heterogeneity in a repetitive DNA element of Fasciola. Int. J. Parasitol., 28, 1923, 1998. 4. Le, T.H. et al. Molecular confirmation that Fasciola gigantica can undertake aberrant migrations in human hosts. J. Clin. Microbiol., 45, 648, 2007.

Fasciola















5. Itagaki, T., and Tsutsumi, K. Triploid form of Fasciola in Japan: genetic relationships between Fasciola hepatica and Fasciola gigantica determined by ITS-2 sequence of nuclear rDNA. Int. J. Parasitol., 28, 777, 1998. 6. Agatsuma, T. et al. Molecular evidence of natural hybridization between Fasciola hepatica and F. gigantica. Parasitol. Int., 49, 231, 2000. 7. Huang, W.Y. et al. Characterisation of Fasciola species from mainland China by ITS-2 ribosomal DNA sequence. Vet. Parasitol., 120, 75, 2004. 8. Itagaki, T. et al. Genetic characterization of parthenogenic Fasciola sp. in Japan on the basis of the sequences of ribosomal and mitochondrial DNA. Parasitology, 131, 679, 2005. 9. Itagaki, T. et al. Molecular characterization of parthenogenic Fasciola sp. in Korea on the basis of DNA sequences of ribosomal ITS1 and mitochondrial NDI gene. J. Vet. Med. Sci., 67, 1115, 2005. 10. Ashrafi, K. et al. Phenotypic analysis of adults of Fasciola hepatica, Fasciola gigantica and intermediate forms from the endemic region of Gilan, Iran. Parasitol. Int., 55, 249, 2006. 11. Lin, R.Q. et al. Sequence analysis of the first internal transcribed spacer of rDNA supports the existence of the intermediate Fasciola between F. hepatica and F. gigantica in mainland China. Parasitol. Res., 101, 813, 2007. 12. Le, T.H. et al. Human fascioliasis and the presence of hybrid/ introgressed forms of Fasciola hepatica and Fasciola gigantica in Vietnam. Int. J. Parasitol., 38, 725, 2008. 13. Urquhart, G.M. et al.. Veterinary Parasitology, Second Edition. Blackwell Science, Oxford, pp. 103–113, 1996. 14. Chen, M.G., and Mott, K.E. Progress in assessment of morbidity due to Fasciola hepatica infection: a review of recent literature. Trop. Dis. Bull., 87, R1, 1990. 15. Mas-Coma, S., and Bargues, M.D. Human liver flukes: a review. Res. Rev. Parasitol., 57, 145, 1997. 16. Mas-Coma, S., Esteban, J.G.., and Bargues, M.D. Epidemiology of human fascioliasis: a review and proposed new classification. Bull. WHO, 77, 340, 1999. 17. Mas-Coma, S. et al., Hepatic trematodiases. In Meyers, W.M., Neafie, R.C., Marty, A.M., Wear, D.J. (Eds.). Pathology of Infectious Diseases, Vol. 1 Helminthiases. Armed Forces Institute of Pathology and American Registry of Pathology, Washington DC, pp. 69–92, 2000. 18. Almendras-Jaramillo, M. et al. Hepatic fascioliasis in children: uncommon clinical manifestations. Arch. Gastroenterol., 34, 241, 1997. 19. Arjona, R. et al. Fascioliasis in developed countries: a review of classic and aberrant forms of the diseases. Medicine (Baltimore), 74, 13, 1995. 20. Kabaalioglu, A., et al. US-guided gallbladder aspiration: a new diagnostic method for biliary fascioliasis. Eur. Radiol., 9, 880, 1999. 21. Prociv, P., Walker, J.C., and Whitby, M. Human ectopic ­fascioliasis in Australia: first case report. Med. J. Aust., 156, 349, 1992. 22. Acosta-Ferreira, W., Vercelli-Retta, J., and Falconi, L.M. Fasciola hepatica human infection. Histopathological study of sixteen cases. Virchows. Arch. A. Pathol. Anat Histol., 383, 319, 1979. 23. Dalimi, A., and Jabarvand, M. Fasciola hepatica in the human eye. Trans. R. Soc. Trop. Med. Hyg., 99, 798, 2005. 24. Zhou, L. et al. Multiple brain hemorrhages and hematomas associated with ectopic fascioliasis in brain and eye. Surg. Neurol., 69, 516, 2008.

793 25. Spithill, T.W., and Dalton, J.P. Progress in development of liver fluke vaccines. Parasitol. Today, 14, 224, 1998. 26. Spithill, T.W., Smooker, P.M., and Copeman, D.B. Fasciola gigantica: epidemiology, control, immunology and molecular biology. In: Dalton, J.P. (Ed.). Fasciolosis. CAB International Publishing, Wallingford, pp. 465–525, 1999. 27. Carrada-Bravo, T. Fascioliasis: diagnosis, epidemiology and treatment. Rev. Gastroenterol. Mex., 68, 135, 2003. 28. Nithiuthai, S. et al. Waterborne zoonotic helminthiases. Vet. Parasitol., 126, 67, 2004. 29. Haseeb, A.N. et al. A review on fascioliasis in Egypt. J. Egypt. Soc. Parasitol., 32, 317, 2002. 30. Ishii, Y., Nakamura-Uchiyama, F., and Nawa, Y. A praziquantelineffective fascioliasis case successfully treated with triclabendazole. Parasitol. Int., 51, 205, 2002. 31. Zhou, P. et al. Food-borne parasitic zoonoses in China: ­perspective for control. Trends Parasitol., 24, 190, 2008. 32. Langley, R.J., and Hillyer, G.V. Detection of circulating immune complexes by the enzyme-linked immunosorbent assay in sera from cattle infected with Fasciola hepatica. J. Parasitol., 75, 690, 1989. 33. Moustafa, N.E., Hegab, M.H., and Hassan, M.M. Role of ELISA in early detection of Fasciola copro-antigens in experimentally infected animals. J. Egypt. Soc. Parasitol., 28, 379, 1998. 34. Sriveny, D. et al. Cathepsin L cysteine proteinase in the diagnosis of bovine Fasciola gigantica infection. Vet. Parasitol., 135, 25, 2006. 35. Maleewong, W. et al. Fasciola gigantica-specific antigens: purification by a continuous-elution method and its evaluation for the diagnosis of human fascioliasis. Am. J. Trop Med. Hyg., 61, 648, 1999. 36. Esteban, J.G.., Bargues, M.D., and Mas-Coma, S. Geographical distribution, diagnosis and treatment of human fascioliasis: a review. Res. Rev. Parasitol., 58, 13, 1998. 37. Hillyer, G..V. Immunodiagnosis of human and animal fasciolosis. In: Dalton, J.P. (Ed.). Fasciolosis. CAB International Publishing, Wallingford, pp. 435–447, 1999. 38. Marcilla, A., Bargues, M.D., and Mas-Coma, S. A PCRRFLP assay for the distinction between Fasciola hepatica and F. gigantica. Mol. Cell. Probes, 16, 327, 2002. 39. Krämer, F. and Schnieder, T. Sequence heterogeneity in a repetitive DNA element of Fasciola. Int. J. Parasitol., 28, 1923, 1998. 40. Mostafa, O.M., Taha, H.A. and Ramadan, G. Diagnosis of Fasciola gigantica in snail using the polymerase chain reaction (PCR) assay. J. Egypt. Soc. Parasitol., 33, 733, 2003. 41. Velusamy, R., Singh, B.P., and Raina, O.K. Detection of Fasciola gigantica infection in snails by polymerase chain reaction. Vet. Parasitol., 120, 85, 2004. 42. Caron, Y., Lasri, S. and Losson, B. Fasciola hepatica: an assessment on the vectorial capacity of Radix labiata and R. balthica commonly found in Belgium. Vet. Parasitol., 149, 95, 2007. 43. Cucher, M.A. et al. PCR diagnosis of Fasciola hepatica in field-collected Lymnaea columella and Lymnaea viatrix snails. Vet. Parasitol., 137, 74, 2006. 44. Magalhães, K.G.. et al. Isolation and detection of Fasciola hepatica DNA in Lymnaea viatrix from formalin-fixed and paraffin-embedded tissues through multiplex-PCR. Vet. Parasitol., 152, 333, 2008. 45. Schweizer, G. et al. Prevalence of Fasciola hepatica in the intermediate host Lymnaea truncatula detected by real time TaqMan PCR in populations from 70 Swiss farms with cattle husbandry. Vet. Parasitol., 150, 164, 2007.

794 46. Luton, K., Walker, D. and Blair, D. Comparisons of ribosomal internal transcribed spacers from two congeneric species of flukes (Platyhelminthes: Trematoda: Digenea). Mol. Biochem. Parasitol., 56, 323, 1992. 47. Adlard, R.D. et al. Comparison of the second internal transcribed spacer (ribosomal DNA) from populations and species of Fasciolidae (Digenea). Int. J. Parasitol., 23, 423, 1993.

Molecular Detection of Foodborne Pathogens 48. Jousson, O. et al. Use of the ITS rDNA for elucidation of some life-cycles of Mesometridae (Trematoda, Digenea). Int. J. Parasitol., 28, 1403, 1998. 49. Hashimoto, K.T. et al. Mitochondrial DNA and nuclear DNA indicate that the Japanese Fasciola species is F. gigantica. Parasitol. Res., 83, 220, 1997.

58 Heterophyidae Ron Dzikowski

The Hebrew University–Hadassah Medical School

Michael G. Levy

North Carolina State University

Contents 58.1. Introduction..................................................................................................................................................................... 795 58.1.1. Fishborne Zoonoses caused by Heterophyidae................................................................................................. 795 58.1.2. Epidemiology..................................................................................................................................................... 796 58.1.3. Clinical Manifestations..................................................................................................................................... 797 58.1.4. Diagnosis........................................................................................................................................................... 798 58.2. Methods........................................................................................................................................................................... 801 58.2.1. Sample Collection and Preparation................................................................................................................... 801 58.2.2. Detection Procedures........................................................................................................................................ 801 58.3. Conclusions and Future Perspective............................................................................................................................... 802 References.................................................................................................................................................................................. 803

58.1  Introduction 58.1.1 Fishborne Zoonoses caused by Heterophyidae Digenea, one of the three major taxa of parasitic Platyhelminthes includes species with a large variety of complex life cycles. They are heteroxenous parasites that require more than one host to complete their life cycle and their adult stages are parasitic in vertebrates. Infections by digeneans are the most common among fishborne zoonoses particularly by species that are less fastidious of their definitive vertebrate hosts. In the past, fishborne zoonotic infections caused by digenea were mainly a problem of either developing countries or of populations that traditionally consumed raw fish. However, global demographic and socio-economical changes together with the development of improved transportation systems have opened global markets for trade and consumption of raw fish products, exposing new populations to the risk of infection. Thus, an increasing number of people are suffering from these zoonoses even in places where the source of infection does not naturally exist.1–2 The current status of fishborne trematode zoonoses has been recently reviewed by several authors.1,3,4 The list of causative agents for human trematodiasis is large and contains liver flukes such as Clonorchis sinesis, Ophistorchis spp. and Metorchis conjunctus as well as intestinal flukes like echinostomes and heterophyids. Species of the family heterophyidae are one example of trematodes that are not fastidious in their definitive or adult stage hosts. They may infect a large variety of vertebrate hosts including piscivorous

birds such as pelicans, herons, and cormorants, as well as a variety of wild and domesticated mammalian species including cats, dogs, and humans. The heterophyidae are very small trematodes: adult worms found in the intestine of their definitive hosts range in length between 0.3 and 3 mm (Figure 58.1). Embryonated eggs, each with a fully developed miracidium are passed in the feces of the host into aquatic habitats where they are ingested by aquatic snails. The miracidia develop in the host snail into sporocysts after which they develop into rediae. Each of the mature rediae may shed hundreds of freeswimming cercariae every day, thus increasing their probability to find and infect fish that serve as the second intermediate hosts. After shedding their tail the cercariae migrate to a specific tissue (muscles, kidney, heart, gills, etc.) encapsulate and develop into a metacercariae (Figure 58.1). Upon ingestion of the infected fish by the final host the metacercariae excyst from their capsule and develop into adult worms that migrate through the digestive tracts to the intestine. Unlike their adult forms which are promiscuous and infect a variety of hosts, larval stages of heterophyids are more specific with regard to their snail and fish hosts. Additionally, most heterophyids exhibits high specificity to the fish tissue in which they will develop to metacercariae. For example, Heterophyes heterophyes, H. dispar, and H. equalis in Middle East develop in the marine snails Pirrenela conica and Cerithidae cingulata, the cercariae infect mainly fish of the families Mugilidae and Cichlidae (tilapia) and migrate to the muscles, where they develop into metacercariae. In endemic regions like the Nile delta in Egypt, where these snails are abundant each fish may harbor hundreds and even thousands of metacercariae. These 795

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Molecular Detection of Foodborne Pathogens

(a)

(b)

(e)

(c)

(f )

(d)

(g)

(h)

50 µm

Figure 58.1  Heterophyid trematodes. (a–c) Adult specimens of Phgicola longa, Haplorchis pumilio, and Dexiogonimus ciureanus collected from Cormorants (Phalacrocorax carbo). (d) Adult specimen of Centrocestus sp. collected from Little Egret (Egretta garzetta). (e) Scanning electron micrograph (SEM) of adult specimen of Pygidiopsis genata collected from Cormorant (P. carbo). (f) Metacercaria of P. genata in the kidney of the cichlid fish Tilapia zillii. (g) Metacercaria of P. longa in the heart of T. zillii. (h) Pleurolophocercous cercaria of Centocestus sp. emerged from the snail Melanoides tuberculata.

minute metacercariae cannot be visually detected without a stereoscope; thus, people consuming the muscles of infected fish will not notice that they are heavily infected. The above reasons may explain why infection with Heterophyes spp. is one of the most common fishborne zoonoses. Different heterophyids develop into metacercariae in different organs. For example, Centrocestus spp. are less fastidious in their choice of fish hosts and infect several fish species including Cichlidae, Cyprinidae, and Poecelidae, and other. They are transmitted by the freshwater snail Melanoides tuberculata and develop into metacercariae in the gills. Other examples for larval migration of heterophyids to specific tissue are Pygidiopsis genata which develops into metacercariae in the kidney; and Phagicola longa, which develops into metacercariae in the heart of freshwater fish such as cichlids.

58.1.2 Epidemiology Foodborne trematode infections drew the attention of the World Health Organization (WHO) as a serious and growing public health concern estimated to affect 40 million people worldwide, whereas the number of people at risk of infection estimated at over half a billion.5–7 In Southeast Asia and the Western Pacific region foodborne trematodiasis is an emerging public health problem.7 It seems that the emergence of infections in these regions is correlated with the exponential growth in aquaculture in endemic regions. Heterophyiasis is not an exception to this trend and is important emerging fishborne zoonoses. Recently, heterophyiasis was documented in regions where it has not been known to be endemic in the past and in some cases their prevalence of infections in those

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Heterophyidae

regions reached remarkably high levels. A recent study in Vietnam has estimated that the prevalence of infection with heterophyiasis in endemic region was 100%. The most abundant species found in human were Haplorchis pumilio and H. taichui.8 In another study cultured fish fom the Mekong Delta, Vietnam were found to be infected with Haplorchis pumilo, H. taichi, Centrocestus formasanus, and Stellantchasmus falcatua with infection rated ranging between 1.7 and 6.6%. Culture conditions as well as seasonal influences were determined to have affected these rates.9 A clinical survey in the southern Philippines, found that 36% of patients that were complaining about abdominal discomfort were suffering from heterophyiasis.10 In a later study these authors reported an exceptionally high prevalence of infection (36%) with H. taichui in human population in another focus in Southern Philippines.11 In Vietnam, in an agricultural province that is populated mainly with farmers with fish ponds integrated into their farming system, a random stool examination demonstrated that the prevalence of infection with small trematode eggs was 65%. The examination of adult worms recovered following anti-helminthic treatment with praziquantel showed that the prevalence of infection with at least one heterophyid species was 100%.8 The dominant species found in these examinations were H. pumilio and H. taichui, and H. yokogawai and Stellanchasmus falcatus were less abundant. The awareness to the epidemiological and clinical significance of various species of intestinal trematodes in the Republic of Korea has recently increased and resulted in several epidemiological studies. In a recent review of those studies, heterophyids, and echinostomes are recognized as the two major groups with significance to public health in terms of both the number of species involved and the number of people infected.12 In this survey, 12 species of heterophyids were documented in humans. The geographical distribution of human infection was found mostly along the eastern and southern coasts with inland endemic foci along certain rivers and streams. The prevalence of infection was found to be between 20 and 70% among the villagers living along these rivers. Surveys conducted at a later time point have demonstrated that new foci of heterophyiasis are emerging,13 mainly by piscivorous birds that migrate between habitats where the snails are thriving. Similarly, high infection rates have been documented in other regions around the globe where heterophyid species are endemic and people are accustomed to eating raw fish. In some regions in Nile delta in Egypt, about a third of the population is infected. The intensity of infection was found to be significantly high among fishermen. This finding is not surprising given the high level of fish consumption, notably grey mullets eaten raw and either dried or mildly salted.14 However, in other places in Egypt where the amount of raw fish consumption is less, the rate of infection was found to be significantly lower. A survey among 3180 patients attending Mansoura University Hospitals’ Clinics, which were subjected to stool examination found that heterophyiasis infection rates were about 4.2%.15 Interestingly, human infection with H. ­heterophyes and H. ­dispar was documented in Korea in people traveling to endemic regions in the Middle

East and were reported as “imported” cases.12 These cases emphasize the need for awareness to ­heterphyiasis as a global concern.

58.1.3 Clinical Manifestations Intestinal heterophyiasis usually cause mild symptoms such as tenderness, chronic diarrhea, bloody diarrhea, abdominal pain, vomiting, etc.4 Symptoms of acid peptic and peptic ulcer disease are also common clinical manifestation of heterophyiasis. The degree of clinical symptoms seems to be related to the intensity of infection: i.e., patients presenting with severe illness tend to have a higher intensity of infection. However, the severity of the symptoms may also be related to the susceptibility as well as the degree of acquired immunity of the individual patient.12 Heavy infections may cause necrosis of the intestinal mucosa that allows the penetration of heterophyid eggs to the mesenteric blood circulation. These eggs may reach different organs and have been reported to cause granulomatous lesions in the myocardium and the brain and spinal cord.4,16 In some cases, patients with heterophyiasis also had cardiac symptoms that improved after anti-­helminthic treatment12 suggesting a possible ectopic parasitism by heterophyids. Recently, experimental infections of H. heterophyes in rats have demonstrated that worms and/or eggs may cross the intestinal wall and spread to visceral organ such as the intestinal wall, lymph nodes, liver, and spleen.17 These results suggest that under certain circumstances, humans—especially immunocompromised patients—may be at risk of ectopic infection with H. heterophyes. The intestinal histopathology caused by heterophyiasis has mainly been determined from studies using experimental infections in animal models such as mice, rats, cats, and dogs. The pathological features were characterized by different degrees of mucosal damage, villous atrophy, hyperplasia, and inflammatory reactions,12 with some evidence for differences in the virulence between species. However, it is extremely difficult to distinguish between the clinical symptoms caused by each individual species in natural infection in humans. This is attributed to the fact that natural infections are usually comprised of multiple species and the lack of speciesspecific diagnostic tools makes it difficult to associate each species with its corresponding symptom(s). In addition, once heterophyid eggs are found in a stool sample, humans are treated with praziquantel which is not species-specific but a general anti-trematode treatment. However, the evidence for ectopic parasitism by some species and the speculation that these species may therefore be more virulent requires a reliable diagnostic tool that would be able to detect heterophyiasis early enough to administer treatment before they cause damage to the central nervous system and without the need for tissue biopsy. In addition, useful diagnostic tools of early stages of heterophyids may have great potential as preventative measures. Early identification of heterophyids in snails and fish may be used by public health authorities to take measures that would prevent infected fish of reaching the markets and would minimize the prevalence of human infections.

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58.1.4  Diagnosis The complex and variable life cycle of digenea is one of the challenging aspects in understanding the biology of these pathogens. Among digenea, heterophyids are among the most difficult to identify at the species level and require special expertise due to their minute size. In addition, precise species determination of early stages such as cercariae emerging from snails and metacercariae infecting fish is ambiguous because they lack the morphological traits of the reproductive system that develop only in the adult worms. Moreover, our knowledge about the complete life cycles is limited to some of the species and experimental demonstrations of life histories are laborious and often unachievable due to the unknown identity of the intermediate and/or definitive hosts. The current methodologies available for accurate diagnosis have serious limitations. The diagnosis of adult heterophyid worms in their definitive host such as dogs, cats, and human is performed by identifying the morphological characteristics of their eggs in stool samples. Staining with potassium permanganate may help to differentiate between eggs of different species, as has been demonstrated for O. viverrini, H. taichui, and Phaneropsolus bonnie.18 However, distinguishing between different species of heterophyid eggs is extremely difficult because of the high similarity in shape and size. Detection of heterophyid eggs in stool sample may be confirmed by recovering the adult flukes after anti-helminthic treatment. Alternatively, serological tests have been used to screen humans for antibodies against parasitic infection.4,12 Several immunodiagnostic tests were evaluated in dogs that were experimentally infected with H. heterophyes.19 These authors concluded that the use of the skin ID test, which is independent of circulating antibodies, may be used for scientific and laboratory diagnoses in naturally infected animals and thus have a potential in the routine diagnosis of Heterophyes infection in humans. However, since the skin ID test did not offer optimal sensitivity and specificity, it is necessary to standardize procedures for antigen preparation and serum testing before firm recommendations can be made. A reported case of human infection with H. nocens was diagnosed by sectional morphology in a biopsy specimen of the small intestine20 however, this rare case obviously does not offer a procedure for routine diagnostic tests. These diagnosis difficulties not only limit our knowledge about the basic biology of these pathogens, but also inhibit disease prevention and treatment. In view of the disease burden of heterophyiasis, the complex life histories of these worms, and the difficulties and limitations of the current diagnosis methodologies, new approaches to develop effective diagnostic tools for these pathogens are essential. Precise identification of the immature stages—­cercariae from snails and metacercariae in fish—is even a greater challenge considering the changes in their morphological traits. Although the morphology changes dramatically during the life cycle of digenea their genetic composition remains the same. This enables targeting of the specific nucleotide sequence of their DNA in order to develop

Molecular Detection of Foodborne Pathogens

genetic markers for precise identification of each species at any point of their life cycle. Additional advantage to this molecular approach is the ability to amplify DNA using polymerase chain reaction (PCR); therefore, only a very small amount of DNA template is required for these reactions. PCR-based diagnostics are therefore very sensitive and allow particular identification of a single cercaria, metacercaria, and adult worm,21 as well as detection of early sporocyst and rediae infections in snails even before they start shedding cercariae.22 Several PCR-based methodologies have been developed and successfully applied for specific diagnostics and molecular taxonomy of fish pathogens (see a comprehensive up to date review by Ted Clark in Ref. 23). As a first step, PCR is used to amplify the gene of interest using gene specific oligonucletide primers. The PCR products are called amplicons and they may be subjected to direct sequencing. An alignment of the DNA sequence of a particular gene can be used to detect species-specific sequence identity or polymorphism. As an alternative to direct sequencing, the amplicons may be used for a PCR-RFLP (restriction fragment length polymorphism) assay. In this assay, the amplicons are subjected to restriction digest that produce sequence-specific cutting patterns based on the affinity of different restriction enzymes to their specific binding sites. RFLP is a faster and more cost effective way to detect polymorphisms, as there is no need for sequence information. However, it may be not as accurate as comparing the sequence itself. Using RFLP, sequence polymorphism can be detected only if it is at the site where the restriction enzyme binds to and cuts the DNA; therefore lack of differences in the restriction patterns may be false negative, if the polymorphisms are found away from the restriction site. It is necessary to perform several restriction digests with different enzymes. However, there is always a possibility that the same digest patterns may still be of different species. Therefore, to increase the accuracy of diagnoses, specific RFLP assay should be designed to differentiate between known sequences that would result in different cutting patterns. Another PCR-based methodology to detect genetic diversity is to use arbitrary primers in order to randomly amplify polymorphic DNA (RAPD). Sequence polymorphism in the genome of different organisms will result in different amplification patterns that can be visualized when separating the amplicons on an agarose gel, creating a typical genetic fingerprint for each organism. RAPD-PCR does not require prior knowledge of the nucleotide sequence and is a useful tool to study population genetics. However, there is evidence that the sensitivity of RAPD-PCR may differentiate between populations of the same species and therefore its suitability for species-specific diagnosis is not clear.24–25 Unveiling species-specific sequence polymorphisms at a certain locus could be used for the development of speciesspecific molecular probes that will specifically anneal to a ­certain sequence. These probes can be used as species-specific primer sets for PCR amplification that will only amplify certain species or as multiplexes of primers that after PCR

Heterophyidae

amplification will result in aplicons of different sizes, each specific for particular species. This method was developed for differentiation of several tremadode species including species of the heterophyidae family.21,22,26,27 In the last 15 years the amount of data of trematodes’ sequence has accumulated rapidly and has been used to study their interrelationships and molecular phylogeny using sequence comparisons.28 The accumulation of the available sequence data was also used to elucidate life histories traits. This became possible by unveiling the genetic linkage through sequence identity between the adult worms that can be identified morphologically and sequenced to the sequence of the same locus in the immature stages. The life histories of few marine and freshwater species have been elucidated using molecular techniques, demonstrating their advantages for specific diagnosis. An early example of the successful use of molecular techniques is the demonstration of the three-host life cycle of Bivesicula claviformis.29 These authors were able to show in an elegant experiment that the mesocercaria found in the gut of the herbivorous fish Thalassoma lunare (Labridae) is genetically identical to and adult worm found in the gut of the carnivore Epinephelus fasciatus (Serranidae). Similar approaches were further used to study life history traits of many other marine species. Anderson was able to link the matacercaria and the adult of Indodidymozoon pearsoni (Digenea: Didymozoidae) by comparing the DNA sequence of fragments of their rDNA.30 Similarly, Jousson, and Bartoli have elucidated the life cycles of some Mesometridae (Trematoda, Digenea)31 and Monorchiidae species.32 Sequence analysis was also used to linked some undescribed larval stages of the digenean family Opecoelidae and the adult worms,33,34 and to distinguish between closely related species of the Opecoelidae family.35 The most common gene targets for PCR amplification in diagnostic tests and phylogenetic studies of trematodes are probably the ribosomal RNA (rRNA) genes (which are commonly referred to as rDNA genes, which represent the underlying sequences in the genome encoding rRNA). The rDNA genes contain three coding regions that encode for the three subunits: the 18S small subunit (SSU), the 5.8S and the 28S large subunit (LSU). The noncoding sequence between the 18S and the 5.8S subunits is the internal transcribed spacer 1 (ITS1); that between the 5.8S and the 28S subunits is the ITS2. The regions of the rDNA that have been mostly used for species differentiation are the 18S small subunit and the ITS regions. The slow evolution rate of these genes in eukaryotes is reflected in the high degree of conservation of their nucleotide sequence. This conservation among species makes them ideal for molecular taxonomy as well as for the development of molecular probes for species-specific diagnostics. Although the ITS regions have been extensively and successfully used to elucidate trematodes life histories29–31 it seems that the ITS regions are more rapidly evolving than the 18S and intraspecific variation has been found in these loci in some trematodes.36,37 Therefore, while it is possible that the use of probes to the ITS regions may be able to distinguish between closely related species, it may also differentiate between strains of the same species. Therefore, it is possible that species-­specific probes

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designed to hybridize to the ITS regions may achieve false negative results, and effectively discriminate between different strains of the same species. Consequently, for speciesspecific diagnosis it is recommended to use the 18S sequence as the main target for comparison, and the ITS regions can be used to confirm the 18S diagnosis or to detect variation among strains of poorly differentiated species. In spite of their importance as foodborne pathogens that affects millions of people worldwide the number of species of heterophyid trematode that have been sequenced is still limited. However, in recent years this data has been increasing constantly (Table 58.1) enabling further development of molecular diagnosis methodologies. Molecular genetics have also used to demonstrate ecological phenomena. In recent studies, the sequences of the ITS1 together with the mitochondrial cytochrome oxidase subunit 1 (mDNA COI) genes were used differentiate cercariae at the species level. Cryptic species of the digeneans, Cercaria batillariae (Heterophyidae) and an undescribed philophthalmid, were identified using PCR-RFLP and sequence analyses of these genes.38 Subsequent sequence comparisons of these cercariae were further used to demonstrate a marked reduction in the genetic variation in one of the species that were introduced to North America, suggesting that cryptic species exhibit different invasion pathways.39 The 28rDNA and mDNA COI sequences were used to confirm that Metagonimus yokogawai, M. takahashii, and M. miyatai are separate species.40 Alignments of heterphyids’ rDNA showed high level of sequence similarity. In fact, a comparison of the 18S sequence of six species of adult worms collected from the guts of piscivorous birds has shown that their base-pair content was 96–98% identical.27 However, the alignment also revealed a polymorphic region in the sequence of the 18S subunit of the gene, about 1400 bp downstream of the start codon, in which all six species differ. This polymorphic region was the target sequence for designating species-specific nucleotide primers for specific PCR amplification of Centrocestus sp. H. ­taichui, H. pumilio, Dexiogonimus ciureanus, P. longa, and P. genata. In order to differentiate between Haplorchis spp. and D. ciureanus it was necessary to perform PCR-RFLP on the entire 18S subunit due the high sequence similarities. Using this diagnosis assay, it was possible to genetically link single cercaria, metacercariae, and adult worms of Centrocestus sp. H. pumilio, P. longa, and P. genata. This study was the first demonstration of the complete life cycle of P. genata. Using this molecular technique, the adult worm collected from cormorants (Phalacrocorax carbo) was genetically linked to metacercaria from the fish Tilapia zillii. It also revealed that the snail Melanopsis costata is the first intermediate host by demonstrating that the cercaria it shed is genetically identical to the adult worm. The successful amplification and sequencing of the rDNA genes of these heterophyids was a prerequisite for the development of the species-specific molecular probes as well as the differentiating PCR-RFLP assay. This procedure has proved itself as a useful, cost effective and above all, an accurate molecular tool for diagnosis of heterophyids that are pathogenic in both fish and human. The high sensitivity of

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Table 58.1 List of Heterophyidae Sequences from GenBank Heterophyid Species Centrocestus sp. Centrocestus formosanus Haplorchis pumilio Haplorchis taichui Haplorchis taichui Phagicola longa Galactosomum lacteum Galactosomum lacteum Pygidiopsis genata Dexiogonimus ciureanus Phagicolinae sp. Cryptocotyle lingua Cryptocotyle lingua Haplorchoides sp. Haplorchoides sp. Heterophyes nocens Heterophyes nocens Metagonimus sp. Metagonimus sp. Metagonimus takahashi Metagonimus takahashi Metagonimus yokogawai Metagonimus yokogawai Metagonimus yokogawai Metagonimus yokogawai Pygidiopsis summa Pygidiopsis summa Stellanchasmus falcatus Stellanchasmus falcatus Stictodora lari

GenBank Accession Number AY245699 AY245759 AY245706 AY245705 EF055885 AY245703 AY222120 AY222227 AY245710 AY245702 AY245766 AJ287492 AJ222228 / AF023111 AJ287521 AY222226 AF188119 AF181890 AF095333 AF096232 AF095332 AF096231 AF095331 AF096230 AF125905/6/7 AF121840/1/2/3/4 AF181885 AF181884 AF181886 AF181887 AY452519 / 20 / 21

these techniques, as well as the fact that the primers amplify only parasites’ DNA and does not cross react either with the snails or the fish DNA, makes them a great diagnostic tool for early detection of possible risk of infection. Applying these molecular tools in aquaculture systems and fisheries in endemic regions may have important practical contribution to global public health interests. In most of these systems, fish are cultured in open ponds where the snail hosts are abundant and piscivorous bird may transmit infection through contaminating the ponds with heterophyid eggs in their feces. In addition to the economical risk that results from fish loss, there is a great risk for initiating heterophyiasis in humans as demonstrated by epidemiological surveys.8 In order to be able to take preventative measures the infection must be detected at early stages of the life cycle. In this regard the complex life cycle of heterophyids is advantageous. DNA extracts from snail samples can be pooled and used as the template for PCR amplification. Using this approach, it is possible to sample snails routinely in order to detect early infections with heterophyid sporocysts and rediae even before they are infective to the fish; i.e., before the snails start shedding ceracariae. Early detection

Sequence Description rDNA (18S-ITS1-5.8S-ITS2) rDNA 18S rDNA (18S-ITS1-5.8S-ITS2) rDNA (18S-ITS1-5.8S-ITS2) mRNA (cytochrome c oxidase subunit I) rDNA (18S-ITS1-5.8S-ITS2) rDNA (18S) rDNA (28S) rDNA (18S-ITS1-5.8S-ITS2) rDNA (18S-ITS1-5.8S-ITS2) rDNA (18S) rDNA (18S) rDNA (28S) rDNA (18S) rDNA (28S) mRNA (cytochrome c oxidase subunit I) rDNA (28S) rDNA (28S) mRNA (cytochrome c oxidase subunit I) rDNA (28S) mRNA (cytochrome c oxidase subunit I) rDNA (28S) mRNA (cytochrome c oxidase subunit I) Cysteine proteinase mRNA Cysteine proteinase mRNA rDNA (28S) mRNA (cytochrome c oxidase subunit I) rDNA (28S) mRNA (cytochrome c oxidase subunit I) Actin gene

of heterophyids in the snails in a certain pond could consequently result in preventative measures, such as transferring the fish and treating the pond against the snails, which will lead to reduce the risk of infection. Similarly, these diagnostic tests may be used by public health agencies to prevent consumption of raw fish infested with heterophyids metacercariae. DNA samples from the muscles and other organs of fish that are known to harbor heterophyids metacercariae, such as grey mullets (Mugilidae) and tilapia (Cichlidae), can be PCR amplified and tested for the presence of heterophyid metacercariae. Early detection of such infection would contribute to the reduction of the risk and the prevalence of infection by classifying these fish products as “risky” so that they are not consumed raw but only if properly cooked or deep frozen prior to consumption. In addition to the PCR-based methods mentioned above that have been successfully applied, other approaches that have been used to diagnose other pathogens can be modified and be used for heterophyiasis diagnostics. For example, solid-phase hybridization techniques like in situ hybridization (ISH) and fluorescence in situ hybridization (FISH)23 may be considered as a useful tool for identification of heterophyids in

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histological samples of snail and fish tissues and in some rare cases from human biopsies. However, at this point it seems that the PCR-based methodologies mentioned above are more convenient for routine diagnosis purposes. Molecular techniques for the diagnosis a number of zoonotic protozoan parasites such as Cryptosporidium, Toxoplasma, Cyclospora, Giardia, and Microsporidia have been developed and are utilized in developed countries. With the exception of Toxoplasma for which methods are available for detection of tissue stages as well as infectious oocysts, these methodologies have primarily focused on detection of infectious cysts contaminating food or water. Infectious cysts are generally present as surface contaminants, and culture methodologies, even when available, are not suitable for the biological amplification of parasite numbers. Therefore, most efforts for detection of protozoan parasites have focused on direct detection of these organisms either directly from food samples of following concentration and purification of infectious cysts (reviewed by Ortega41). In contrast to various techniques developed for concentrating cystic forms of protozoan parasites that may be utilized in conjunction with molecular assays, manual removal of these minute heterophyid trematodes from tissue samples is laborious and therefore unlikely to be of value for the screening of fresh fish.

58.2  Methods In this chapter we will focus on the methodologies that have been already proved to be successful for the diagnostics of species of heterophyid species. We will give detailed explanations of the techniques one can use to collect the genetic material, extract DNA from different organisms and tissues and to amplify the gene of interest by PCR in order to be able to sequence new species. We will also outline the basic rules for designing species-specific primer sets and multiplexes for specific diagnosis. We hope that this chapter will provide knowledge that would benefit anyone that wants to use similar approach to expand the available molecular tool box for heterophyid diagnosis.

58.2.1 Sample Collection and Preparation Development of specific PCR assays have been developed for a large number of trematode species, including heterophyids.22,27,28 Adults trematodes, that can be identified to species based on internal morphology are collected by sedimentation of the intestine content of the final host (piscivorous birds, cats, dogs, mice or rats) then removed to clean physiological saline and preserved in 70% ethanol. Metcercariae of heterophyids are collected directly from the tissue of the infected fish (gills, subcutaneous, heart, conus arteriosus, kidney, liver). They should be removed gently from the surrounding tissue by needles under a stereoscope and preserved in 70% ethanol. Cercariae are collected from aquatic snails by exposing the infected snails to strong light. And then individual specimens can be either frozen at -70°C or fixed in 70% ethanol (see Table 58.2).

Table 58.2 List of Reagents and Equipment Required Reagents Sterile distilled water 10 × PCR buffer MgCl2 dNTPs Primer sets Taq polymerase Aagarose Gel electrophoresis buffer Ethidium bromide

Equipment DNA thermal cycler Gel electrophoresis apparatus Electrophoresis power supply UV transilluminator Sterile, disposable plastic microtubes

To isolate DNA, approximately 25 mg of tissue should be dissected from tissue of choice and placed into a sterile microcentrifuge tube. Lysis of the tissue is achieved by incubating at 56°C in 180 μl Qiagen buffer ATL plus 20 μl Qiagen proteinase K. Incubation can vary from 1 h to overnight in order to ensure complete lysis. Following lysis, the Qiagen DNEasy Tissue kit should be utilized following manufacturer protocol for DNA purification.

58.2.2  Detection Procedures It is preferable to use freshly isolated or specimens preserved by freezing or in 70% alcohol rather than in solutions such as formalin that make recovery of DNA difficult. rDNA genes are often the target of such efforts as the presence of multiple copies of these genes in the genome generally increases the sensitivity of the assays as well as a large number of published gene sequences useful for development of oligonucleotide primers for initial sequencing. Once these genes are sequenced, they are aligned with similar genes from related species and unique sequences identified. Oligonucleitide primer sets are designed that complement these species-specific regions and are generally designed to bracket approximately a 150–1000 bp segment of the target gene. Specific PCR assays for fish-infesting trematodes have generally utilized conventional PCR assays in which the final products are manually transferred to agarose or a similar gel matrix, electrophoresed, and the resulting DNA amplicons, if present, visualized following staining with a flurochrome such as ethidium bromide whose fluorescence when excited with ultraviolet light increases approximately 20 fold when intercolated into double stranded DNA. The use of real-time PCR (RT-PCR) which incorporates a flurochrome directly into the PCR reaction mixture which is detected as the reaction progresses eliminates the need for post-PCR processing and provides for significant saving of time as well as reduces the possibility of laboratory contamination with target DNA as reaction tubes need not be opened while increasing sample throughput.42 Although it is not yet possible to connect all stages of these parasites based on morphology, as previously stated the DNA is constant between life cycle stages. Thus once a PCR assay with the desired degree of specificity

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has been developed, it may be applied to the detection and identification of any stage of the parasite.21,22,27,29–31 Molecular detection of trematodes in fish flesh will require extraction of DNA from representative samples of fish tissue. Following release of DNA which requires one to several hours of proteinase K digestion, DNA is purified utilizing one of several available methods and the presence of heterophyid-specific DNA determined. This in a time-consuming process, generally requiring greater than 24 h from the time samples are received in the laboratory and cost constraints will likely mean that limited numbers of samples can be processed (i) PCR amplification: PCR amplification can be done on DNA extracted from a pool of trematodes or from individual worms, metacercariae, and cercariae. Specimens should be washed overnight in 10 mM TrisEDTA buffer. Following another two washes (1 h each), a single metacercariae or cercaria is placed directly into a 0.2 ml PCR tube and gently crushed, prior to DNA extractions or direct amplification. DNA is extracted from single adult warms using DNeasy tissue kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. PCR amplification is carried out in 20–50 µl volumes using 10 × buffer, 2.5 U Taq polymerase, 10 mM of each deoxynucleotide triphosphate, and 100 ng of each primer. The PCR conditions should be optimizing for each of the primers that are used and according to the size of the expected amplicons. PCR amplicons are subjected to electrophoresis in 1.5% agarose gels, stained with ethidium bromide and visualized under ultraviolet light. PCR products can be purified using commercial PCR purification kit and sequenced directly. As an alternative the amplicons of interest can be cut out of the gels, purified, and cloned into pGEM-T Easy Vector system, utilizing blue/white selection. Plasmids from white colonies should be screened for inserts using PCR amplification. Positive colonies are incubated overnight and then the plasmids are purified and sequenced. Procedure: (1) Remove reagents from freezer to thaw. (2) Place a 1.5-ml microcentrifuge tube, and enough PCR tubes for all samples into a clean area, preferably into a hood specifically designed for molecular applications that allows for decontamination via ultraviolet light. (3) Once reagents are thawed and tubes exposed to UV, prepare a Master Mix in sufficient volume for all samples. Include at least two extra tubes to be used as positive and negative controls. (4) To a sterile 1.5-ml microcentrifuge tube, add 37.5 μl sterile H2O, 5 μl 10 × PCR buffer, 3 μl MgCl2, 1 μl each of forward and reverse primers, and 0.5 μl Taq polymerase for each sample to be run. (5) Mix well by pipetting several times, then aliquot 49 μl of the Master Mix into each PCR tube. (6) To each sample tube, add 1 μl purified sample DNA.

Molecular Detection of Foodborne Pathogens

(7) To the positive control tube add 1 μl known positive DNA. (8) Add 1 μl sterile water or known negative sample to the negative control. (9) Close tubes, place into thermal cycler and start necessary program. (10) After completion of PCR program, run the samples, controls, and appropriate DNA size marker on a 1.5% agarose gel containing 1 μl of a 1% solution of ethidium bromide per 100 ml agarose until sufficiently separated. (11) Visualize results by placing the gel onto a UV transilluminator. (12) DNA bands should be analyzed by comparing to the DNA size markers. Where the sizes are appropriate, a positive diagnosis can be made on proven assays, or the bands can be sequenced for confirmation. (ii) Endonuclease digestion: PCR-RFLP procedure can be performed using a 5 µl aliquot of the rDNA 18S PCR product (1.5 µg) of each species and digest it with restriction enzymes according to the manufacturer’s specification, at 37ºC for 4 h. Electrophoresis of the resulting fragments is done on a 1.5% agarose gel and visualized with ethidium bromide. (iii) Interpretation: Interpretation of PCR results may however be problematic. Positive reactions are generally considered to be indicative of the presence of the suspected organism, but this is not always the case. Besides indicating a true positive, such an outcome may also result from the presence of contamination with DNA from another sample, laboratory contamination and/or. contamination with cross reacting but “nontarget species” DNA should the PCR assay not be sufficiently specific. Careful sample handling from collection through final processing is required because parasite DNA is readily transferred due to the highly stable nature of DNA and the extreme sensitivity of PCR assays may detect minute amounts of such contaminants. Similarly, negative reactions may indicate a true negative, but may also result from insufficient target material in the sample tested, DNA degradation or the presence of inhibitors of the PCR assay.

58.3 Conclusions and Future Perspective The advances in molecular techniques in the last two decades as well as the increase in the amount of the genetic data available in open accessed computerized databases such as GenBank has set the stage for the development of new diagnostic tools for pathogens. The field of molecular diagnosis of pathogen is evolving rapidly and new techniques are being developed for precise, cost effective, and sometimes high throughput diagnostics. These advances have been used to promote the development of molecular diagnostic tools for precise identification of many causative agents of zoonoses, including

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Heterophyidae

fishborne pathogens. The worldwide emergence of fishborne zoonoses emphasizes the need for accurate diagnostic tools to determine the best course of clinical treatment for patients. In addition, the complexities of the life history of treamotodes can indeed be advantageous by providing an opportunity for detection of infection foci of early stages heterophyid even before they become infective. Therefore, the great potential of molecular diagnosis is that it would give health authorities a great monitoring tool, which is precise, inexpensive, and with the right training, simple to use. Upon early detection of infection, these tools will allow to execute preventative measures to be executed in order to limit the scope of infection. In spite of its great potential, the development of molecular diagnostic tools for heterophyidae is still at its very early stage. It is surprising that although heterophyiasis is a disease that affects millions worldwide, the volume of molecular research of this particular trematode family is still low. As a consequence, the available sequence data, which is the basis for the development of molecular diagnostic tools, is still limited. Nevertheless, the methodologies that have been developed and applied successfully to sequence and specifically diagnose some of the species described in this chapter can be, and hopefully will be applied for other heterophyid species. In order to develop these tools researchers will first have to overcome the lack of sequence data of heterophiyidae by sequencing the species of interest. Previously the amplification of the rDNA from species lacking prior sequence data required the use of oligonucleotide primers that have been designated to anneal to the conserved regions flanking the gene of interest-in most cases the 18S subunit of the rDNA gene. These conserved regions have been found by alignment of known sequence of many other trematode species. However, some of the conserved primers that successfully amplified most trematode rDNA genes failed to amplify properly the heterophyids rDNA (Dzikowski, unpublished data). Thus, sequencing was possible only after purification and sub-cloning of the PCR fragments into pGEM-T vector. The accumulation of rDNA sequence data of heterophyids should help to generate new alignments and allow the design of better primers for more efficient amplification and sequencing of additional heterophyid species. The foundations for the development of species-specific molecular probes set up in the work reviewed here, will hopefully contribute to the expansion of these tool and to develop diagnostic tools for other heterophyid species that are important disease causing agents in humans, such as H. heterophyes, H. dispar, H. nocens, Pygidiopsis stanma, and others. It will be possible in the near future to have molecular diagnostic tools for each of the heterophyid species. However, in order to reach that goal a global effort in research and development will have to take place to collect specimen, sequence them and to develop molecular diagnostic assays. Expansion of the available database of heterophyid sequences will allow the development of more complex PCR-based methodologies that are sensitive enough to differentiate between species through the detection of single nucleotide polymorphism (SNPs) such as PCR/LDR43 and high resolution melt (HRM).44 Because of the high sequence similarity among the heterophyidae, these techniques may

prove themselves in the future as the ultimate diagnostic tool for infections. For example using PCR/LDR methodology, a multiplex of species-specific primer sets can be designed that can differentiate between sequences with only one base-pair difference. In the future one can envision a “heterophyidae multiplex” of primers that will give precise identification for each individual species in a single reaction. In a recent report of the WHO, it was recommended that governments in countries where fishborne trematodiasis is endemic would implement control programs into the existing health care system, continue surveillance measures, and practice to eliminate the risk of infection from aquaculture systems and products.6 These control procedures would greatly benefit from having good molecular diagnostic tools for identification of larval stages of heterophyids in snails and fish in fisheries and aquaculture systems. Health care authorities will be able to accurately monitor the prevalence of infection among humans, snails, and fish. Early detection of infection foci will allow the implementation of preventative steps to protect the human population from heterophyiasis.

References

1. Chai, J.Y., Murrell, K.D. and Lymbery, A.J. Fish-borne ­parasitic zoonoses: status and issues. Int. J. Parasitol., 35, 1233, 2005. 2. Macpherson, C.N. Human behaviour and the epidemiology of parasitic zoonoses. Int. J. Parasitol., 35: 1319, 2005. 3. Fried, B., Graczyk, T.K. and Tamang, L. Food-borne ­intestinal trematodiases in humans. Parasitol. Res., 93: 159, 2004. 4. Ko, R.C. Fish-borne parasitic zoonoses. In Fish Diseases and Disorders, Vol. 1. Woo, P.T.K. (ed.), pp. 592–628. CAB International, Cambridge, MA, 2006. 5. WHO. Foodborne Trematode Infections. WHO Tech. Rep. Ser. No. 849, pp. 1–157, Geneva, Switzerland, 1995. 6. WHO. Foodborne Trematode Infections in Asia, p. 55. World Health Organization, Manila, Phillipines, 2004. 7. Keiser, J. and Utzinger, J. Emerging foodborne trematodiasis. Emerg. Infect. Dis., 11, 1507, 2005. 8. Trung Dung D. et al. Fishborne zoonotic intestinal trematodes. Vietnam. Emerg. Infect. Dis., 13, 1828, 2007. 9. Thien, S.U. et al. Prevalence of fishborne zoonotic parasites in important cultured fish species in the Mekong Delta, Vietnem. Parasitol Res., 5, 1277, 2007. 10. Belizario, V.Y., Jr. et al. Intestinal heterophyidiasis: an emerging food-borne parasitic zoonosis in southern Philippines. Southeast Asian J. Trop. Med. Public Health, 32 (Suppl 2), 36, 2001. 11. Belizario, V.Y. et al. A focus of human infection by Haplorchis taichui (Trematoda: Heterophyidae) in the southern Philippines. J. Parasitol., 90, 1165, 2004. 12. Chai, J.-Y., and Lee, S.-H. Food-borne intestinal trematode infections in the Republic of Korea. Parasitol. Int., 51, 129, 2002. 13. Park, J.H. et al. A new endemic focus of Heterophyes nocens and other heterophyid infections in a coastal area of Gangjingun, Jeollanam-do. Korean J. Parasitol., 45, 33, 2007. 14. Abou-Basha, L.M. et al. Epidemiological study of heterophyiasis among humans in an area of Egypt. East Mediterr. Health J., 6, 932, 2000. 15. El Shazly, A.M. et al. Intestinal parasites in Dakahlia governorate, with different techniques in diagnosing protozoa. J. Egypt Soc. Parasitol., 36, 1023, 2006.

804 16. Africa, C.M., de Leon, W., and Garcia, E.Y. Visceral complications in intestinal heterophyidiasis of man. Acta Med. Philippina, Monographic Series, 1, 1, 1940. 17. Elsheikha, H.M. Heterophyosis: Risk of ectopic infection. Vet. Parasitol., 147, 341, 2007. 18. Sukontason, K., Piangjai, S. and Chaithong, U. Potassium ­permanganate staining for differentiation the surface morphology of Opisthorchis viverrini, Haplorchis taichui and Phaneropsolus bonnei eggs. Southeast Asian J. Trop. Med. Public Health, 30, 371, 1999. 19. Elshazly, A.M. et al. Comparison of three immunodiagnostic tests for experimental Heterophyes heterophyes infection in dogs. Vet. Parasitol., 151, 196, 2008. 20. Ryang, Y.S. et al. An incidental case of human Heterophyes nocens infection diagnosed by sectional morphology in a biopsy specimen of the small intestine. Korean J. Parasitol., 37, 189, 1999. 21. Dzikowski, R. et al. Clinostomum complanatum and Clinostomum marginatum (Rudolphi, 1819) (Digenea: Clinostomidae) are separate species based on differences in ribosomal DNA. J. Parasitol., 90, 413, 2004. 22. Levy, M.G. et al. Morphologic, pathologic, and genetic investigations of Bolbophorus species affecting cultured channel catfish in the Mississippi delta. J. Aquatic Anim. Health, 14, 235, 2002. 23. Clark, T.K. Molecular approaches and techniques. In Fish Diseases and Disorders, Vol. 1 Protozoan and Metazoan Infections. Woo, P.T.K. (ed.), p. 725–752. CAB International, Cambridge, MA, 2006. 24. Cunningham, C.O. and Mo, T.A. Random amplified polymorphic DNA (RAPD) analysis of three Norwegian Gyrodactylus salaris populations (Monogenea: Gyrodactylidae). J. Parasitol., 83, 311, 1997. 25. Martin-Sanchez, J. et al. Molecular arguments for considering Hysterothylacium fabri (Nematoda: Anisakidae) a complex of sibling species. Parasitol. Res., 89: 214, 2003. 26. Dzikowski, R. et al. Genetic and morphologic differentiation of Bolbophorus confusus and B. levantinus (Digenea: Diplostomatidae), based on rDNA SSU polymorphism and SEM. Dis. Aquatic Organisms, 57, 231, 2003. 27. Dzikowski, R. et al. Use of rDNA polymorphism for identification of heterophyidae infecting freshwater fishes. Dis. Aquatic Organisms, 59, 35, 2004. 28. Cribb, T.H.B. et al. The Digenea. Interrelationship of the Platyhelminthes. Littlewood, D.T.J.B. (ed.), p. 168–185. Taylor & Francis Inc., London, 2001. 29. Cribb, T.H. et al. A DNA-based demonstration of a three-host life-cycle for the Bivesiculidae (Platyhelminthes: Digenea). Int. J. Parasitol., 28, 1791, 1998.

Molecular Detection of Foodborne Pathogens 30. Anderson, G.R. Identification and maturation of the metacercaria of Indodidymozoon pearsoni. J. Helminthol., 73, 21, 1999. 31. Jousson, O. et al. Use of the ITS rDNA for elucidation of some life-cycles of Mesometridae (Trematoda, Digenea). Int. J. Parasitol., 28, 1403, 1998. 32. Bartoli, P., Jousson, O. and Russell-Pinto, F. The life cycle of Monorchis parvus (Digenea: Monorchiidae) demonstrated by developmental and molecular data. J. Parasitol., 86, 479, 2000. 33. Jousson, O., Bartoli, P. and Pawlowski, J. Molecular identification of developmental stages in Opecoelidae (Digenea). Int. J. Parasitol., 29, 1853, 1999. 34. Jousson, O. and Bartoli, P. The life cycle of Opecoeloides ­columbellae (Pagenstecher, 1863) n. comb. (Digenea, opecoelidae): evidence from molecules and morphology. Int. J. Parasitol., 30, 747, 2000. 35. Jousson, O. and Bartoli, P. Molecules, morphology and morphometrics of Cainocreadium labracis and Cainocreadium dentecis n. sp. (Digenea: Opecoelidae) parasitic in marine fishes. Int. J. Parasitol., 31, 706, 2001. 36. Sorensen, R.E., Curtis, J. and Minchella, D.J. Intraspecific variation in the rDNA its loci of 37-collar-spined echinostomes from North America: implications for sequence-based diagnoses and phylogenetics. J. Parasitol., 84, 992, 1998. 37. van Herwerden, L., Blair, D. and Agatsuma, T. Intra- and interindividual variation in ITS1 of Paragonimus westermani (Trematoda: digenea) and related species: implications for phylogenetic studies. Mol. Phylogenet. Evol., 12, 67, 1999. 38. Miura, O. et al. Molecular-genetic analyses reveal cryptic species of trematodes in the intertidal gastropod, Batillaria cumingi (Crosse). Int. J. Parasitol., 35, 793, 2005. 39. Miura, O. et al. Introduced cryptic species of parasites exhibit different invasion pathways. Proc. Natl. Acad. Sci. USA, 103, 19818, 2006. 40. Lee, S.U. et al. Sequence comparisons of 28S ribosomal DNA and mitochondrial cytochrome C oxidase subunit I of Metagonimus yokagawai, M. takahashii and M. miyatai. Korean J. Parasitol., 42, 129, 2004. 41. Ortega, Y.R. Molecular tools for the identification of foodborne parasites. In PCR Methods in Food, Maurer, J. (ed.), pp. 119–145. Springer, New York, NY, 2005. 42. Heid, C.A. et al. Real time quantitative PCR. Genome Res., 10, 986, 1996. 43. Favis, R. and Barany, F. Mutation detection in K-ras, BRCA1, BRCA2, and p53 using PCR/LDR and a universal DNA microarray. Ann. N. Y. Acad. Sci., 906, 39, 2000. 44. Liew, M. et al. Genotyping of single-nucleotide polymorphisms by high-resolution melting of small amplicons. Clin. Chem., 50, 1156, 2004.

59 Metagonimus Jae-Ran Yu

Konkuk University School of Medicine

Jong-Yil Chai

Seoul National University College of Medicine

Contents 59.1 Introduction.................................................................................................................................................................... 805 59.1.1 Classification and Morphology of the Genus Metagonimus............................................................................. 806 59.1.1.1 Metagonimus yokogawai (Katsurada, 1912).................................................................................... 806 59.1.1.2 Metagonimus takahashii (Suzuki, 1930)......................................................................................... 806 59.1.1.3 Metagonimus miyatai (Saito, Chai, Kim, Lee, and Rim, 1997)...................................................... 806 59.1.2 Biology, Pathogenesis, and Medical Importance.............................................................................................. 806 59.1.2.1 Biology............................................................................................................................................. 806 59.1.2.2 Pathogenicity.................................................................................................................................... 806 59.1.2.3 Medical Importance......................................................................................................................... 807 59.1.3 Identification and Diagnosis............................................................................................................................. 807 59.2 Methods.......................................................................................................................................................................... 808 59.2.1 Reagents and Equipment................................................................................................................................... 808 59.2.2 Sample Collection and Preparation.................................................................................................................. 808 59.2.3 Detection Procedures........................................................................................................................................ 808 59.2.3.1 PCR-RFLP....................................................................................................................................... 808 59.2.3.2 SSR-PCR.......................................................................................................................................... 809 59.2.3.3 RAPD . ............................................................................................................................................ 809 59.2.3.4 Sequencing of 28S D1 rDNA and mtCOI PCR............................................................................... 809 59.2.3.5 Karyotyping......................................................................................................................................810 59.3 Conclusion and Future Perspectives................................................................................................................................810 References...................................................................................................................................................................................810

59.1  Introduction Trematodes of the genus Metagonimus are intestinal flukes of humans and animals, having a small or minute body, and they belong to the family Heterophyidae (heterophyid flukes). The genus Metagonimus was created by Katsurada in 1912 based on adult flukes recovered from experimental dogs given the flesh of sweetfish, Plecoglossus altivelis, in Taiwan, and also from people who consumed raw sweetfish.1 At present, four species, namely M. yokogawai, M. takahashii, M. miyatai, and M. minutus, are known to infect humans.2 Metagonimiasis, an infection caused by one or more species of Metagonimus, is prevalent as an important human disease in the Far East, including Republic of Korea, China, Japan, and far eastern Russia.2 The diagnosis is usually based on recovery of eggs in the feces of patients. However, it is necessary at times, for example, in light infection cases, to use immunological and molecular techniques. Metagonimus species flukes have a small body size (1.0– 1.5 mm in length) with a laterally deviated ventral sucker,

and lack the ventrogenital apparatus or genital sucker.2 They are morphologically differed from other heterophyid flukes such as Heterophyes, Heterophyopsis, Pygidiopsis, Centrocestus, Stellantchasmus, Stictodora, Haplorchis, and Procerovum in the following points. Heterophyes and Heterophyopsis differ from Metagonimus in having a bigger, median located ventral sucker and a prominent genital sucker.3 Pygidiopsis has a median located ventral sucker connected anterolaterally with a ventrogenital apparatus.3 Centrocestus has a median located ventral sucker and small spines on the oral sucker.3 Stellantchasmus has a small, laterally deviated ventral sucker, as in Metagonimus, but differs in the presence of an elongated sac-like seminal vesicle with a muscular expulsor at the opposite side of the ventral sucker.3 Stictodora, Haplorchis, and Procerovum have an armed gonotyl superimposed on the laterally deviated ventral sucker.3 Haplorchis and Procerovum have only one testis, whereas Metagonimus and other heterophyids have two testes.3 805

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59.1.1 Classification and Morphology of the Genus Metagonimus In the genus Metagonimus, total six species have been reported;2 M. yokogawai Katsurada, 1912, M. takahashii Suzuki, 1930, M. minutus Katsuta, 1932, M. katsuradai Izumi, 1935, M. otsurui Saito and Shimizu, 1968, and M. miyatai Saito et al., 1997. Among them, four species, excluding M. katsuradai and M. otsurui, have been reported from human infections.2 59.1.1.1  Metagonimus yokogawai (Katsurada, 1912) This species was originally described in Japan, and is the most prevalent of the three species of Metagonimus. This species is now known to be distributed in Japan, Korea, China, Taiwan, and Russia.3 The characteristic morphological features include the presence of laterally deviated ventral sucker, without a genital sucker, two testes adjacent to each other near the posterior end of the body, and vitelline follicles distributed laterally between the level of the ovary and middle portion of the posterior testis. The adult worms measure 1.0–2.0 mm long and 0.4–0.6 mm wide. Eggs measure 28–30 mm long and 16–17 mm wide. 59.1.1.2  Metagonimus takahashii (Suzuki, 1930) This species was first reported in Japan, by Takahashi in 1929, in the small intestine of mice and dogs fed Metagonimus metacercariae encysted in several species of freshwater fish not including the sweetfish; 1 year later it was described as a new species.4 It differs from M. yokogawai in the position of two testes (anterior testis separated from the posterior testis), the distribution of vitelline follicles (more abundant and crossing over the posterior-most end), and by larger size of eggs (M. yokogawai, 28–30 mm; M. takahashii, 32–36 mm).4 The Koga type of Metagonimus encysting in the dace Tribolodon spp. is regarded a synonym of M. takahashii.5 Human infection was suggested when eggs were detected in feces in 1988.6 Adult flukes were first confirmed in 1993 in inhabitants of Umsong-gun, Chunchungnam-do, along the upper reaches of the Namhan River.5 59.1.1.3 Metagonimus miyatai (Saito, Chai, Kim, Lee, and Rim, 1997) This species was first reported in Japan in 1941, but its taxonomic significance was questioned.7 Thus, it was designated as the “Metagonimus Miyata type.”7 In 1997, its validity was acknowledged and described as a new species.7 The description was based on adult flukes collected from dogs and hamsters experimentally fed the metacercariae encysted in the sweetfish, dace, common fat-minnow Morocco steindachneri, pale chub Zacco platypus, dark chub Zacco temmincki, and also on specimens collected from naturally infected humans.7 This species is morphologically different from M. yokogawai and M. takahashii in the position of the posterior testis (separated greatly from the anterior one), the distribution

Molecular Detection of Foodborne Pathogens

of vitelline follicles (never crossing over the posterior testis), and the intermediate size of eggs (28–32 μm).7 In a study on numerical taxonomy, M. miyatai (under the name of the Miyata type) was suggested to be a subspecies of M. takahashii.8 However, M. miyatai is genetically distinct from M. takahashii.9–11 Metagonimus miyatai eggs are larger than the eggs of M. yokogawai but smaller than those of M. takahashii, and the adult flukes recovered were morphologically close but incompatible with M. yokogawai or M. takahashii.12 A high prevalence of this fluke infection was reported among the people residing around the river.13–16

59.1.2 Biology, Pathogenesis, and Medical Importance 59.1.2.1 Biology Metagonimus yokogawai (Katsurada, 1912). The molluscan first intermediate host is a kind of the fresh water snail, Semisulcospira coreana or S. libertina.17 The most important second intermediate host is the sweetfish P. altivelis,2 but the dace Tribolodon sp.5 and the perch Lateolabrax japonicus18 also serve as the second intermediate hosts. Dogs, rats, and cats were reported as natural definitive hosts,19–21 although their significance as the source of the human infection (i.e., as reservoir hosts) has not been established. Metagonimus takahashii (Suzuki, 1930). The source of metacercariae is the crucian carp Carassius carassius.22 Human infection was suggested when eggs were detected in feces in 1988.6 Adult flukes were first confirmed in 1993 in inhabitants of Umsong-gun, Chunchungnam-do, along the upper reaches of the Namhan River.13 The snail host involved may be Semisulcospira coreana or Koreanomelania nodifila,17 but this has not yet been established. The fish hosts are the crussian carp,22 the carp Cyprinus carpio,7 and the dace Tribolodon taczanowskii.5 There are no reports on the reservoir host. Metagonimus miyatai (Saito, Chai, Kim, Lee, and Rim, 1997). Semisulcospira globus14 was reported to be the first intermediate host. Freshwater fish, Zacco platypus and Zacco temminckii, are the second intermediate hosts.2 Mice,12 rats,5,14 hamsters,5,7 and dogs14,23 are experimental definitive hosts. The reservoir host is unknown. 59.1.2.2  Pathogenicity The adult flukes of M. yokogawai are found to parasitize in the middle part of the small intestines; within the crypt of Lieberkühn in the early stages of the infection (by day 2–3, postinfection), and between the villi in the later stages.24–26 The pathological features are characterized by villous atrophy and crypt hyperplasia, with variable degrees of inflammatory reactions. The infected mucosa show blunting and fusion of the villi, edema of the villus tips, congestion, and inflammatory cell infiltrations in the villous stroma, and decreased

Metagonimus

villus/crypt height ratios. In immunocompetent animals, the location of the worms is confined to the intestinal mucosa.26–28 However, immunosuppression of ICR mice by prednisolone injection allow a deeper invasion of the worms into the submucosa or underneath.29 Immunosuppression enhances the survival of the worms and prolongs their life spans in the same mouse strain.29,30 Similar histopathological features are seen in the small intestines of M. miyatai-infected mice, but the degree of the mucosal damage is less severe than in M. yokogawai-infected mice, as represented by the stronger expression patterns of the proliferating cell nuclear antigen (PCNA) in the intestinal mucosa.31 In immunocompetent mice, the intestinal histopathological changes due to M. yokogawai infection are normalized at 3–4 weeks after the infection.30 59.1.2.3  Medical Importance In patients infected with M. yokogawa, the most frequent symptoms reported are mild to moderate degrees of abdominal pain, diarrhea, lethargy, anorexia, and weight loss.17,32 The degree of clinical symptoms seems to be related to the individual worm burdens; heavier infection cases tend to suffer from severer illnesses. However, the severity of the symptoms may also be related to the susceptibility as well as the degree of acquired immunity of the individual patient. A new visitor to an endemic area, for example, suffered from a severe illness after a primary infection,33 whereas long-term residents in endemic areas generally complained of milder symptoms than those expected.32 The drug of choice for all infections by heterophyids is praziquantel. The efficacy of a single oral dose of 10–20 mg/kg praziquantel is satisfactory; for example, up to a 95–100% cure rate for M. yokogawai infection.34,35

59.1.3 Identification and Diagnosis Eggs of M. yokogawai can be differentiated from other heterophyid eggs by their length of 26.9–31.6 mm, elliptical shape with length/width ratio of 1.5–2.1, clean shell surface, less prominent operculum, no shoulder rims, and dark yellow or brown color.36 The eggs of M. takahashii and M. miyatai have similar morphology with those of M. yokogawai, with the exception of the larger egg sizes of the two former species; hence, the measurement of egg size is essential. There could be false egg negative cases among the light infection cases with M. yokogawai, for example, with less than 100 worms in an infected person. The number of eggs produced per day per worm for M. yokogawai was reported to be only 14–64 eggs in the human host,16 so the detectability of eggs in feces from such a case is negligible. Serological tests such as ELISA are helpful in false egg negative cases.33,37 There are several studies supporting strong evidences of three Metagonimus species are genetically different. The three species of Metagonimus are genetically differentiated by the polymerase chain reaction-based restriction

807

fragment length polymorphism (PCR-RFLP) patterns.9 Internal transcribed spacer 1 (ITS1) site of ribosomal RNA and mitochondrial cytochrome c oxidase I (mCOI) gene were targeted for PCR-RFLP. The restriction patterns of ITS1 gene with RsaI, AluI, and MspI showed multiple fragmented bands of different sizes between three kinds of Metagonimus.9 Especially in mCOI gene, restriction pattern with RsaI and AluI produced differentially fragmented band patterns.9 However, they did not provide gene sequence of ITS1 and mCOI genes of three species of Metagonimus. PCR-based random amplification of polymorphic DNA (RAPD) technique using random 10-mer oligonucleotide primers also showed distinguishable banding patterns between M. yokogawai and M. miyatai.38 Previous studies regarding the number of chromosomes and karyotypes of genus Metagonimus supported strong evidence of genetic differences between M. yokogawai and M. miyatai.10 They showed that the number of bivalents in the first meiotic division of M. takahashii was nine (n=9) and the diploid number of M. miyatai and M. yokogawai were 18 (2n=18) and 32 (2n=32).10 In addition, chromosomes of M. miyatai were consisted of one pair of metacentric, seven pairs of submetacentric, and one pair of telocentric chromosomes and those of M. yokogawai were two pairs of metacentric, 11 pairs of submetacentric, and three pairs of subtelocentric chromosomes.10 Simple sequence repeat anchored polymerase chain reaction using primers of 3′ or 5′ termini of the (CA)n repeats showed distinguishable banding patterns among the three species of Metagonimus, suggesting that they have different genotypes.11 The study on sequence comparison of 28S ribosomal DNA and mCOI subunit I of three Metagonimus species also supported M. miyatai as a separate species.39 The length of the 28S D1 region rDNA sequence of the three Metagonimus species is 248 bp and its G+C content is 52%.39 The length of the mtCOI sequence averaged 400 bp with a G+C content ranged from 44% (M. miyatai), 46% (M. takahashii) to 47% (M. yokogawai).39 Nucleotide sequence differences between species were 23.0% (92/400 bp) between M. miyatai and M. takahashii, 16.2% (65/400 bp) between M. miyatai and M. yokogawai, 13.2% (53/400 bp) between M. takahashii and M. yokogawai.39 Nucleotide gaps sequence differences were 2.5% (10/400 bp) between M. miyatai and M. takahashii, 2.7% (11/400 bp) between M. miyatai and M. yokogawai, and 2.4% (11/400 bp) between M. takahashii and M. yokogawai.39 The aligned sequences of three Metagonimus species showed high similarities with other comparative human intestinal trematodes (P. summa and S. falcatus) for 28S D1 rDNA (95.0% and 94%) and the mtCOI gene (68.5% and 78.0%).39 M. takahashii and M. yokogawai placed in the same clade supported by DNA sequence and phylogenetic tree analysis for 28S D1 rDNA and mtCOI gene by neighbor-joining and parsimony method. M. miyatai placed in different clade from two other Metagonimus species.39

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59.2  Methods 59.2.1 Reagents and Equipment (1) DNA extraction: lysis buffer (10 mM Tris-Cl pH 8.0, 100 mM EDTA, 0.5% SDS), proteinase K, RNase A, phenol, chloroform, isoamyl alcohol, 5 M NaCl, 99% ethanol, TE buffer (10 mM Tris-Cl pH 8.0, 1 mM EDTA). (2) PCR: PCR buffer, MgCl2, dNTP, primers, Thermus aquaticus DNA polymerase, DNA template. (3) Electrophoresis: agarose LE gel, TAE buffer, ­bis-acrylamide, ethidium bromide, silver staining solution. (4) Southern blot: restriction enzymes RsaI, AluI MspI, CfoI, and TaqI; DIG nucleic acid detection kit. (5) PCR cloning and sequencing: primera, ExTaq enzyme, EcoRV site of a pT7Blue T-vector kit plus ligase, NovaBlue competent cell, IPTG, X-gal, BamHI, and HindIII enzymes. (6) Reagents: 1× liquid MEM-199 medium, colchicines, Carnoy’s solution (one part of glacial acetic acid, three parts of 95% ethanol), 45% acetic acid, acetic­ orcein solution, Canada balsam solution. (7) Supplies: Eppendorf tube, gavage needle, pipette, liquid nitrogen, mortar, and pestle, nylon membrane, RESISITE, CALCD, UPGMA. QIAEX DNA gel extraction kit, QIAprep spin plasmid kit, Sequenase kit, automated DNA sequencer, dissecting pin, forceps, slide glass, cover glass, alcohol lamp, light microscope. (8) Other materials: NCBI databases, EMBOSS GEECEE program, CLUSTAL W program, TREEVIEW tree drawing software, NeighborJoining, phylogeny inference package (PHYLIP). (9) Equipment: 37oC incubator, centrifuge, PCR thermocycler, SDS-gel caster. (10) Fish and animals: Plecoglossus altivelis, Zacco platypus, Carassius carassius, SD rat

59.2.2 Sample Collection and Preparation Sample collection: Metacercariae of M. yokogawai are collected by artificial digestion of P. altivelis caught at Oshipcheon (stream), Samchok-gun, Kangwon-do. Metacercariae of M. miyatai (formerly Metagonimus miyatai type) and M. takahashii are collected from Z. platypus and C. carassius caught at Talchongang (River), Chungju, Chungchongbuk-do. Three kinds of adult worms are obtained from SD rats through oral infection. Collected worms are stored at –20oC until being used for DNA extraction. DNA extraction (1) Pulverize the adult worms of three kinds using a LN-cooled mortar and pestle. (2) Place a 1/10 volume of pulverized tissue in a 1.5-ml tube containing lysis buffer and incubated overnight at 37oC.

Molecular Detection of Foodborne Pathogens

(3) Add proteinase K (100 μg/ml), RNase A (20 μg/ml) and incubate for 2 h at 37oC. (4) Add an equal volume of phenol: chloroform: isoamyl alcohol and mix by inverting. (5) Precipitate total genomic DNA overnight with addition of 1/20 volume 5 M NaCl and 2 volume 99% ethanol at –70oC. (6) Centrifuge at 10,000×g for 10 min. (7) Resuspend in 50 μl of TE buffer. (8) Add 0.1 volume of RNase A (10 mg/ml) and incubate for 1 h at 37oC. (9) Remove residual phenol by chloroform extraction. Resuspend the genomic DNA in 25 μl of TE buffer. (10) Store at 4oC.

59.2.3  Detection Procedures 59.2.3.1  PCR-RFLP Principle: We present a PCR-RFLP analysis for ­discrimination among the three species of Metagonimus, M. yakogawai, M. takahashii, and M. miyatai.9 The conserved ITS1 sequence between 18S rRNA and 5.8S rRNA genes is amplified by PCR with primers BD1 (5′-GTCGTAACAAGGTTCCGTA-3′) and 4S (5′-TCTAGATGCGTTCGAAGTGTCGATG-3′). The ITS1 probe (including 5.8s rRNA) is prepared with primers 4S and 3S (5′-GGTACCGGTGGATCACTCGGCTCG-3′) and labeled with DIG labeling kit. The mitochondrial COI is produced with primers JB3 (5′-TTTTTTGGGCATCCTGAGTTTAT-3′) and JB4.5 (5′-TAAAGAAAGAACATAATGAAAATG-3′). The ITS1 and mitochondrial COI fragments are digested with restriction enzymes RsaI, AluI, MspI, CfoI, and TaqI, separated on 3% agarose gel, transferred to nylon membrane and probed with DIG labeled ITS1 probe. Southern blotting is done as follows.9 PCR products of BD1 and 4S are electrophoresed through 1% agarose LE gels (Tris-acetate EDTA). DNA fragments are transferred onto a nylon membrane which is probed by digoxigenin-labeled 5.8S rRNA gene. DIG nucleic acid detection kit is used to detect the bands. Procedure: (1) Prepare PCR mixture (20 μl) containing: • 1× PCR buffer, 2.5 mM MgCl2, 100 μM dNTP, 2.5 U of Taq DNA polymerase. • 25 pmol of each primer pair (BD1 and 4S for ITS1, 4S, and 3S for ITS1 probe, or JB3 and JB4.5 for mitochondrial COI). • 1 μl (90 ng) of template DNA, and distilled water to 20 μl. (2) Perform PCR amplification in a thermocycler with the following cycling program: for ITS1: 45 cycles of 95oC for 45 sec, 51oC for 1 min, and 72oC for 90 sec. For mCOI: 45 cycles of 94oC for 1 min, 50oC for 1 min, and 72oC for 2 min. (3) Use DIG probe synthesis kit to making ITS1 probe (5.8S rRNA gene detecting ITS1).

809

Metagonimus

Note: The results of ITS1 and mitochondrial COI digestion are expected to reveal unique banding patterns among the three species of Metagonimus. In fact, based on PCR-RFLP patterns of ITS1 and mitochondrial COI genes, it has been suggested that M. miyatai evolved at older times than the other two species of Metagonimus.9 59.2.3.2 SSR-PCR Principle: A simple sequence repeat-PCR (SSR-PCR) is also useful for comparison and differentiation among three Metagonimus species.11 Metacercariae of M. yokogawai, M. miyatai, and M. takahashii are collected from freshwater fish, P. altivelis, Z. platypus, and C. carassius, respectively, caught at different localities. Metacercariae are isolated using an artificial digestion technique. Three kinds of adult worms are obtained from SD rats through oral infection and sacrifice on day 14 postinfection. Collected worms are stored at –70oC until genomic DNA extraction using the method outlined in Section 59.2.2. SSR-PCR is performed with genomic DNA from the three Metagonimus species using 3′ or 5′ termini of (CA)n repeated primers (see below). Procedure: (1) Prepare PCR mixture (20 μl) containing: • 1× PCR buffer, 2.5 mM MgCl2, 100 μM dNTP, 2.5 U of Taq DNA polymerase. • 0.5 μM of each of the following primers with 3′ or 5′ termini of (CA)n repeats: −− three kinds of 3′ anchored primers: (CA)8RG, (CA)4RG, and (CA)8RY (all at 0.5 μM/μl) −− two kinds of 5′ anchored primers: BDB(CA)7C, DBDA(CA)7 (all at 0.5 μM/μl) • 1 μl (20 ng) of template DNA, and distilled water to 20 μl. (2) Perform PCR amplification in a thermocycler with the following cycling program: 30 cycles of 94oC for 30 sec, 52oC for 45 sec and 72oC for 2 min. (3) Separate the amplified products on 1% agarose gel and detect by ethidium bromdide staining. Note: Although SSR-PCR fingerprinting for genomic DNA generally demonstrates a higher level of polymorphism than PCR-RFLP or RAPD analyses, it can be applied to determine the phylogenic relations of three species of Metagonimus.11 The differences of SSR-PCR patterns among the three species might be related to their morphological differences and biological differences, for example, in taking up different fish intermediate hosts.11 59.2.3.3 RAPD Principle: RAPD analysis can be applied to distinguish adult flukes of M. yokogawai and M. miyatai.38 Procedure: (1) Prepare PCR mixture (20 μl) containing: • 1× PCR buffer, 2.5 mM MgCl2, 100 μM dNTP, 2.5 U of Taq DNA polymerase.

• 25 pmol of each of the following random primer (in separate tube): OPA-02 (5′-TGCCGAGCT G-3′), OPA-09 (5′-GGGTAACGCC-3′), OPA-10 (5′-GTGATCGCAG-3′), OPA-11 (5′-CAATCGC CGT-3′), OPA-13(5′-CAGCACCCAC-3′), OPA-17 (5′-GACCGCTTGT-3′), OPA-19 (5′-CAAACGT CGG-3′), or OPA-20 (5′-GTTGCGATCC-3′). • 1 μl (90 ng) of template DNA, and distilled water to 20 μl. (2) Perform PCR amplification in a thermocycler with the following cycling program: 45 cycles of 94oC for 1 min, 35oC for 1 min and 72oC for 2 min. (3) Separate the amplified products on 8% polyacrylamide gel and detect by silver staining. Note: The above eight primers were proved to be useful for differentiating M. yokogawai from M. miyatai, which have differential morphology and biology including the kinds of fish hosts involved.38 59.2.3.4 Sequencing of 28S D1 rDNA and mtCOI PCR Principle: Comparison of sequences of 28S D1 ribosomal DNA (28S D1 rDNA) and mtCOI fragments can be utilized to differentiate among the three species of Metagonimus: M. yokogawai, M. takahashii, and M. miyatai.39 Adult specimens of M. takahashii, M. miyatai, and M. yokogawai are obtained from SD rats experimentally infected with metacercariae at day 7 postinfection. Metacercariae of each species are collected from freshwater fish, C. carassius, Z. platypus, and P. altivelis, respectively. Two species of the family Heterophyidae, Pygidiopsis summa and Stellantchasmus falcatus, are obtained from experimentally infected animals, and S. falcatus is used as an outgroup to infer phylogeny. Procedure: (1) Prepare PCR mixture (20 μl) for D1 rDNA and mtCOI PCR containing: • 1× PCR buffer, 2.5 mM MgCl2, 100 μM dNTP, 2.5 U of ExTaq enzyme. • 25 pmol of each of the following random primer: 28S rDNA: JB10 (forward), 5′-GATTACCCGC TGAACTTAAGCATA-3′. JB9 (reverse), 5′-GCTGCATTCACAAACACC CCCACTC-3′. mtCOI: JB3 (forward), 5′-TTTTTTGGGCA TCCTGAGGTTTAT-3′ (2575). JB4.5 (reverse), 5′-TAAAGAAAGAACATAA TGAAAATG-3′ (3021) (primers from the nucleotide sequences of Fasciola hepatica). • 1 μl (10 ng) of template DNA, and distilled water to 20 μl. (2) Perform PCR amplification in a thermocycler with the following cycling program: 40 cycles of denaturation at 95oC for 20 sec, annealing at 55oC for 30 sec (for 28S rDNA) or 48oC 30 sec (for mtCOI),

810

extension at 72oC for 30 sec; and 1 cycle of final extension at 72oC for 6 min. (3) Sequence the PCR products. Note: The sequences of 28S rDNA and mitochondrial COI genes are useful for discrimination among the three species of Metagonimus.39 The phylogenetic trees established by 28S rDNA and mitochondrial COI nucleotide sequences revealed that M. yokogawai is more closely related with M. takahashii than with M. miyatai, which supports the results of PCRRFLP.9 59.2.3.5 Karyotyping Principle: Karyotyping of M. yokogawai, M. takahashii, and M. miyatai is performed to analyze the chromosome numbers and karyotypes.10 Adult specimens of M. takahashii, M. miyatai, and M. yokogawai are obtained from SD rats at day 7 postinfection. Metacercariae of each species are collected from C. carassius, Z. platypus, and P. altivelis, respectively. Procedure: (1) Cultivate worms in 1× liquid MEM-199 medium containing 0.001% colchicines, at 37oC for 4 h. (2) Fix with modified Carnoy’s fluid. (3) Remove the testes of each specimen with a dissecting pin and transfer to the fixative using a fine forceps. (4) Remove pieces of the other tissues. (5) Mince the testes gently in 45% acetic acid to prepare cell suspensions. (6) Stain with aceticorcein solution for 10–30 min. (7) Rinse briefly overstained cell with 45% acetic acid and cover with cover glass, and then squash. (8) Heat the slide briefly with alcohol lamp to remove air bubbles. (9) Mount in Canada balsam. (10) Observe under a light microscope; measure the relative length of chromosomes, total chromosome length of the haploid set, and the positions of centromeres. Note: The diploid chromosome number of M. yokogawai is shown to be 32 (2n = 32), whereas that of M. miyatai was 18 (2n = 18).10 Karyotyping of M. takahashii may be difficult, as it failed in a study.10

Molecular Detection of Foodborne Pathogens

the perch, Lateolabrax japonicus. Humans are contracted by these flukes by consuming raw or improperly cooked flesh of these fishes. When infected, the flukes parasitize the small intestine and cause mucosal inflammation with villous atrophy and crypt hyperplasia. The infected patients may experience episodes of diarrhea and abdominal discomfort, together with anorexia and weight loss. The diagnosis is based on recovery of eggs from the feces. It can be treated with praziquantel. For the purpose of taxonomic researches and also for use as a diagnostic aid, molecular techniques have been applied. Molecular techniques applied in studying Metagonimus spp. include PCR-RFLP, SSR-PCR, RAPD, sequencing of 28S rDNA and mtCOI, and karyotyping. Based on the results of molecular studies, M. yokogawai and M. takahashii may be phylogenetically close to each other. M. miyatai is considered to have evolved at an earlier time than M. yokogawai or M. takahashii. Further studies are required to draw firm conclusions regarding the phylogenetic relationships among the species of Metagonimus.

References





59.3 Conclusion and Future Perspectives



Human metagonimiasis caused by M. yokogawai, M. takahashii, M. miyatai, and M. minutus is prevalent in endemic areas of the Republic of Korea, China, Japan, and Russia. In the Republic of Korea, an estimated about 240,000 infected cases are present along the eastern and southern coastal areas.2 The first intermediate host of M. yokogawai is the freshwater snail, Semisulcospira libertina or S. coreana. The second intermediate hosts include freshwater fishes, including the sweetfish, P. altivelis, the dace, Tribolodon sp., and





1. Ito, J. Metagonimus and other human hererophyid trematodes. Progr. Med. Parasitol. Jpn., 1, 314, 1964. 2. Chai, J.Y. and Lee, S.H. Food-borne intestinal trematode infections in the Republic of Korea. Parasitol. Int., 51, 129, 2002. 3. Chai, J.Y. Intestinal flukes. In Food-borne Parasitic Zoonoses: Fish and Plant-borne Parasites, Murrell, K.D. and Fried, B. (Eds.). Springer, New York, 2007. 4. Suzuki, S. Yokogawa’s Metagonimus. List of publications on special animals in Okayama prefecture. Okayama Prefectural Report, 146, 1930. 5. Chai, J.Y. et al. Three morphological types of the genus Metagonimus encysted in the dace, Tribolodon taczanowskii, caught from the Sumjin River. Korean J. Parasitol., 29, 217, 1991. 6. Ahn, Y.K. Epidemiological studies on Metagonimus yokogawai infection in Samcheok-gun, Kangwon-do, Korea. Korean J. Parasitol., 22, 161, 1984. 7. Saito, S. et al. Metagonimus miyatai sp. nov. (Digenea: Heterophyidae), a new intestinal trematode transmitted by freshwater fishes in Japan and Korea. Korean J. Parasitol., 35, 223, 1997. 8. Kim, K.H. et al. A numerical taxonomic study on heterophyid trematodes. Korean J. Parasitol., 29, 55, 1991. 9. Yu, J.R. et al. PCR-RFLP pattern of three kinds of Metagonimus in Korea. Korean J. Parasitol., 35, 271, 1997. 10. Lee, S.U. et al. A cytogenetic study on human intestinal trematodes of the genus Metagonimus (Digenea: Heterophyidae) in Korea. Korean J. Parasitol., 37, 237, 1999. 11. Yang, H.J. et al. Molecular differentiation of three species of Metagonimus by simple sequence repeat anchored polymerase chain reaction (SSR-PCR) amplification. J. Parasitol., 86, 1170, 2000. 12. Kim, C.H. Study on the Metagonimus sp. in Gum river basin, Chungchung-nam Do, Korea. Korean J. Parasitol., 18, 215, 1980. 13. Chai, J.Y. et al. An epidemiological study of metagonimiasis along the upper reaches of the Namhan River. Korean J. Parasitol., 31, 99, 1993.

Metagonimus 14. Kim, C.H. et al. Studies on the Metagonimus fluke in the Daecheong reservoir and the upper stream of Geum river, Korea. Korean J. Parasitol., 25, 69, 1987. 15. Park, M.S. et al. Intestinal parasite infections in the inhabitants along the Hantan River, Chorwon-gun. Korean J. Parasitol., 31, 375, 1993. 16. Ahn, Y.K. Intestinal flukes of genus Metagonimus and their second intermediate hosts in Kangwon-do. Korean J. Parasitol., 31, 331, 1993. 17. Cho, S.Y., Kang, S.Y. and Lee, J.B. Metagonimiasis in Korea. Arzneimittelforschung, 34, 1211, 1984. 18. Ahn, Y.K. Lateolaborax japonicus, a role of the second intermediate host of Metagonimus yokogawai. New Med. J., 26, 135, 1983. 19. Cho, S.Y., Kang, S.Y. and Ryang, Y.S. Helminthes infections in the small intestine of stray dogs in Ejungbu City, Kyunggi Do, Korea. Korean J. Parasitol., 19, 55, 1981. 20. Seo, B.S. et al. Studies on parasitic helminths of Korea 5. Survey on intestinal trematodes of house rats. Korean J. Parasitol., 19, 131, 1981. 21. Huh, S., Sohn, W.M. and Chai, J.Y. Intestinal parasites of cats purchased in Seoul. Korean J. Parasitol, 31, 371, 1993. 22. Chun, S.K. A study on the metacercaria of Metagonimus takahashii and Exorchis oviformis from Carassius carassius. Bull. Pusan Fish. Coll., 3, 31, 1960. 23. Ahn, Y.K. and Ryang, Y.S. Growth, egg-laying and life span of Miyata type of genus Metagonimus (Trematoda: Heterop­ hyidae) in the final hosts. J. Wonju Med. Coll., 8, 8, 1995. 24. Chai, J.Y. Study on Metagonimus yokogawai (Katsurada, 1912) in Korea V. Intestinal pathology in experimentally infected albino rats. Seoul J. Med., 20, 104, 1979. 25. Lee, J.B. et al. Study on the pathology of metagonimiasis in experimentally infected cat intestine. Korean J. Parasitol., 19, 109, 1981. 26. Kang, S.Y. et al. A study on intestinal lesions of experimentally reinfected dogs with Metagonimus yokogawai. Korean J. Parasitol., 21, 58, 1983. 27. Rho, I.H. et al. Observation on the pathogenesis of villous changes in early phase of experimental metagonimiasis. Chung-Ang J. Med., 9, 67, 1984.

811 28. Jang, Y.K. et al. In situ posture of anterior body of Metagonimus yokogawai in experimentally infected dog. Korean J. Parasitol., 23, 203, 1985. 29. Chai, J.Y., Seo, B.S. and Lee, S.H. Study on Metagonimus yokogawai (Katsurada, 1912) in Korea VII. Susceptibility of various strains of mice to Metagonimus infection and effect of prednisolone. Korean J. Parasitol., 22, 153, 1984. 30. Chai, J.Y., Kim, J. and Lee, S.H. Invasion of Metagonimus yokogawai into the submucosal layer of the small intestine of immunosuppressed mice. Korean J. Parasitol., 33, 313, 1995. 31. Yu, J.R., Myong, N. and Chai, J.Y. Expression patterns of proliferating cell nuclear antigen in the small intestine of mice infected with Metagonimus yokogawai and Metagonimus Miyata type. Korean J. Parasitol., 35, 239, 1997. 32. Seo, B.S. et al. Intensity of Metagonimus yokogawai infection among inhabitants in Tamjin River basin with reference to its eff laying capacity in human host. Seoul J. Med., 26, 207, 1985. 33. Chai, J.Y. et al. An egg-negative patient of acute metagonimiasis diagnosed serologically by ELISA. Seoul J. Med., 30, 139, 1989. 34. Rim, H.J. et al. Anthelmintic effects of various drugs against metagonimiasis. Korean J. Parasitol., 16, 117, 1978. 35. Lee, S.H. et al. Anthelmintic effects of praziquantel (Distocide@) against Metagonimus yokogawai infection. J. Korean Soc. Chemother., 2, 167, 1984. 36. Lee, S.H. et al. Comparative morphology of eggs of heterophyids and Clonorchis sinensis causing human infections in Korea. Korean J. Parasitol., 22, 171, 1984. 37. Cho, S.Y., Kim, S.I. and Kang, S.Y. Specific IgG antibody responses in experimental cat metagonimiasis. Korean J. Parasitol., 25, 149, 1987. 38. Yu, J.R., Chung, J.S. and Chai, J.Y. Different RAPD patterns between Metagonimus yokogawai and Metagonimus miyatai type. Korean J. Parasitol., 35, 295, 1997. 39. Lee, S.U. et al. Sequence comparisons of 28S ribosomal DNA and mitochondrial cytochrome c oxidase subunit I of Metagonimus yokogawai, M. takahashii and M. miyatai. Korean J. Parasitol., 42, 129, 2004.

60 Opisthorchis

Paiboon Sithithaworn and Thewarach Laha Khon Kaen University

Ross H. Andrews

University of South Australia

Contents 60.1 Introduction.................................................................................................................................................................... 813 60.1.1 Classification..................................................................................................................................................... 813 60.1.2 Geographical Distribution.................................................................................................................................814 60.1.3 Biology and Life Cycle......................................................................................................................................814 60.1.4 Pathogenesis and Pathology.............................................................................................................................. 815 60.1.5 Host Immune Response.....................................................................................................................................816 60.1.6 Clinical Manifestation and Morbidity...............................................................................................................816 60.1.7 Liver Fluke Induced Cholangiocarcinoma........................................................................................................817 60.1.8 Epidemiology, Prevention, and Control.............................................................................................................819 60.1.9 Laboratory Diagnosis.........................................................................................................................................819 60.1.9.1 Parasitological Techniques................................................................................................................819 60.1.9.2 Immunological Techniques.............................................................................................................. 820 60.1.9.3 Molecular Techniques...................................................................................................................... 821 60.2 Methods.......................................................................................................................................................................... 821 60.2.1 Preparation of O. viverrini DNA using a Commercial Kit............................................................................... 821 60.2.2 Preparation of O. viverrini DNA using a CTAB Method................................................................................. 822 60.2.3 PCR Detection of O. viverrini from Stool Samples.......................................................................................... 822 60.2.3.1 Use of pOV-6 Primers...................................................................................................................... 822 60.2.3.2 Use of Trem25-F and OV25-4R Primers......................................................................................... 823 60.3 Conclusions and Future Perspectives............................................................................................................................. 823 References.................................................................................................................................................................................. 823

60.1  Introduction “The genus Opisthorchis contains several common liver flukes (e.g., O. viverrini, O. felineus and Clonorchis sinensis (oriental or Chinese liver fluke) (see Chapter 55) in the family Opisthorchiidae.” Opisthorchis viverrini is a carcinogenic fishborne trematode which inhabits the biliary tract. It is an overlooked parasite because it is viewed as a local problem with limited occurrence and because comprehensive epidemiological studies and the extent of morbidity, particularly in southeast Asia, is not fully realized. Recent estimates of populations at risk have indicated that a vast number of people in Asia and the rest of the world are also at risk of infection from consumption of aquaculture imports. In addition to the economic and food safety impact of O. viverrini infection (opisthorchiasis), a significant public health concern is its association with bile duct cancer, cholangiocarcinoma (CCA). CCA is a devastating malignancy and notoriously difficult to diagnose and is associated with high mortality. Unlike other cancers, CCA is in theory

preventable if exposure to the causative agent, O. viverrini, is avoided. Thus accurate and reliable diagnosis of opisthorchiasis is desirable not only for laboratory diagnosis for individuals but also surveillance and control of the parasite as well as lessening the risk of CCA.

60.1.1 Classification Liver flukes are parasites classified in the phylum platyhelminthes (flat worm), class trematoda (fluke), subclass digenea, and order opisthorchiformes. The liver flukes comprise three families, namely; Opisthorchiidae, Fasiolidae, and Dicrocoeliidae. The most commonly reported members as causative agents for human infection are in the family Opisthorchiidae consisting of Opisthorchis viverrini, O. felineus and Clonorchis sinensis. Other less common members in this family are O. conjunctus, O. guayaquilensis, O. noverca, Metorchis albidus, M. comjunctus, Pseudamphistomum aethiopicum, and P. truncatum. Only O. conjunctus has been reported in man while the rest are found predominantly in animal hosts.1 813

814

In the family Fasciolidae and Dicrocoeliidae, four species of liver flukes commonly found in herbivores but which also infect humans are Fasciola hepatica, F. gigantica, Dicrocoelium dendriticum and D. hospes. The generic name Opisthorchis was described in 1875 and Clonorchis in 1907, based on the morphology of adult worms, the major differences separating them were the shape and position of testes and appearance of vitelline glands.2,3 In the case of C. sinensis (see Chapter 58), the adult worm has two highly branched testes situated tandemly and the vitelline glands are distributed continuously along both sides of the body. O. viverrini and O. felineus share similar features by having lobular testes and vitelline glands arranged in clusters. O. viverrini has distinctive and deeper lobulation of testes and lack of a transverse arrangement of vitelline follicles in comparison with O. felineus.

60.1.2 Geographical Distribution Among the three major human liver flukes, the current estimated total number of infections is 45 million in Asia and Europe, of which 9 million people are infected with O. viverrini, 1.2 million with O. felineus and 35 million with C. sinensis.4,5 As many as 680 million people worldwide are at risk of infection by the three liver flukes. The hot spot of O. viverrini is in northeast Thailand, Lao PDR, Cambodia, and to a lesser extent, Vietnam. As many as 67 million people in southeast Asia are estimated to be at risk of infection.6 C. sinensis is found in Korea, China, Taiwan, Vietnam, and previously in Japan while O. felineus is endemic in Poland, Germany, Russia, Federation of Kazakhstan and western Siberia. Being similar to other snailborne trematodes, the presence of suitable snail intermediate host dictates the geographical distributions of these liver flukes. However, to date, because of population migration and extensive traveling plus growing importation of freshwater aquaculture products from Asia, it is now increasingly common to detect infected people in nonendemic areas. For example, cases of opisthorchiasis in nonendemic areas such as the United States and Europe have been reported in immigrants from Asia and the cause of infection was due to the consumption of imported raw or undercooked freshwater fish containing metacercariae.7,8 Similar to most parasitic heminths, high geographical variability has been observed in the distribution of O. viverrini in a given endemic area. For example, in Thailand O. viverrini is highly prevalent in the northeast in contrast to other regions of the country with lower prevalence.9 In Lao PDR, O. viverrini has been found mainly in the southern region such as Saravane, Savannakhet, and Khammouan.10 Very little information is available on the distribution of O. viverrini in Cambodia.11,12

60.1.3  Biology and Life Cycle O. viverrini is a hermaphroditic, dorso-ventrally flattened and leaf like trematode. The body is armed with two muscular suckers, the oral sucker situated anteriorly and the ventral sucker at the mid-body. The size of adult flukes

Molecular Detection of Foodborne Pathogens

measures 5.5–10×0.77–1.65 mm (C. sinensis is larger measuring 10–25×3–5 mm, O. felineus is the smallest measuring 7–12×2–3 mm).3 The size of adult worms may vary depending on the intensity of infection and the diameters of the bile ducts they inhabit. Adult worms inhabit the biliary tract mainly, intrahepatic bile ducts, gall bladder and to a lesser extent the pancreatic duct.13 After development to sexual maturity, the adult worms cross-fertilize and produce yellow-brown eggs that are discharged with bile fluid into the intestine and the feces and passed out into the environment. The eggs measure 25–35 µm in length and 15–17 µm in width; all three species of liver fluke eggs have distinct opercular shoulders surrounding the operculum at one end and a small knob or comma shape appendage at the abopercular end. The egg shell surface is rough and irregular or seen as musk-melon patterns by electron microscopy.14,15 Eggs of the three species of liver flukes share similar morphologies and are difficult to differentiate. The number of eggs produced per worm depends on the worm burden, i.e., a density-dependent effect and pathological condition of the hepatobiliary system. In humans each worm on average produces about 50 egg/g feces.13,16 When eggs reach a body of freshwater (i.e., small ponds, streams, and rivers, flooded rice fields) and are ingested by an appropriate snail, miracidium hatch and develop into sporocysts and rediae. Different species of Bithynia (i.e., B. goniomphalos, B. siamensis siamensis, B. funiculata) serve as the intermediate hosts.2 The rediae gave rise to cercariae which are released daily approximately 2 months after snail infection. Once exposed to appropriate fish intermediate hosts, the free-swimming cercariae shed their tails, penetrate into the tissues or the skin of freshwater fish and become fully infective metacercariae in 3 weeks. Humans become infected by ingesting the metacercariae in uncooked fish. The ingested metacercariae excyst in the duodenum, enter the common bile duct and migrate to the distal bile duct. Development of juveniles in humans requires approximately 2 months to become the sexually mature stage. In the case of the closely related liver fluke, C. sinensis, it has been reported that they can survive up to 26 years in a human host.17 This observation may represent an extreme survival condition in an individual host. Since there has been no direct estimate of the average life expectancy of O. viverrini, it may be assumed that the average survivorship could be shorter. Based on the pattern of the age-intensity profile, it is anticipated that O. viverrini may survive in humans for approximately 10 years. Based on systematic analyses, there is an evidence that natural population of O. viverrini in southeast Asia is a species complex in which adult worms show identical and indistinguishable morphologies but they are genetically very different (i.e., cryptic species). Currently results indicate that at least two cryptic species, one from northeast Thailand and another from Lao PDR occur within the complex.18,19 The supporting genetic evidence for the presence of cryptic species in O. viverrini arise from applications of allozyme electrophoresis and random amplified polymorphic DNA and the association of distinct genetic groups of O. viverrini with wetland

815

Opisthorchis

river systems as well as correlations with specific genetically distinct geographical isolates of Bithynia snails.19 Preliminary evidence of genetic variation for C. sinensis has also been documented in which isolates from China and Korea have shown relatively low degrees of genetic difference similar to the difference detected between C. sinensis and O. viverrini based on nuclear ribosomal DNA and mitochondrial DNA sequences.20,21 To date it is obvious that information on the systematics and population genetics of O. viverrini, C. sinensis as well as O. felineus is limited because a lack of suitable independent genetic markers. Compared to the limited number of genes available currently for genetic and molecular characterization, results from the recent application of the technique of allozyme elecetrophoresis have established up to 32 independent genetic markers that can now be used to examine the systematics and population structure of O. viverrini.19 Similar to other trematodes, the prevalence of O. viverrini infection in snail intermediate hosts (Bithynia sp) is typically less than 1%.22 The density of snail populations exhibit strong seasonality and is dependent on rainfall, being highly abundant in the rainy season and distributed extensively in shallow water and paddy fields; but disappearing rapidly in the dry season. Because of such high fluctuations in numbers, parasite control via control of snail populations does not appear to be a feasible approach. In contrast to the infection in snails, the prevalence of infection in fish intermediate hosts is much higher. As high as 90–95% of several species of cyprinid fish have been reported to harbor O. viverrini metacercariae.23 The most common species of cyprinid fish are in the genera Puntius, Cyclocheilichthys, and Hampala.2 Host finding mechanisms of cercariae is a complex process24 and free swimming cercariae are very efficient in locating the appropriate species of fish in a large volume of water. The intensity of liver fluke infections in fish varies by season, species, individuals, and types of water bodies.23,25 Most metacercariae are distributed throughout the body of fish and some are found in the head of the fish. The metacercarial burden peaks in winter (around October–February) and becomes low in the rainy season and summer thus transmission of the parasite from fish to humans is probably seasonal.26 The number of metacercariae reported in fish varies within a range of between one to hundreds. Frequency distribution patterns of metacercariae of O. viverrini in fish is highly dispersed; most fish have few metacercariae while a few fish harbor a heavy metacercarial load.27 Recent studies which have found trematode metacercariae other than O. viverrini in cyprinid fish in Thailand28,29 have indicated that the occurrence of mixed species of trematodes in a given fish species is common. Careful morphological identification of metacercariae from fish requires experience and trained personnel. Dogs and cats are reservoir hosts in the life cycle of the liver fluke but the prevalence of infection in these animals is relatively low.30 Although fecal contamination from infected animals undoubtedly contributes to parasite transmission to snails in the environment, its actual importance is unclear.

However, in a situation where fecal contamination from humans is reduced or eliminated by mass treatment and with improved sanitation, infection among reservoir hosts may serve to maintain the source of parasite to play a critical role in transmission and viability of the life cycle.

60.1.4  Pathogenesis and Pathology The pathological changes in liver fluke infection are confined mainly to bile ducts, the liver, and gall bladder in both human and animal models. The magnitude of the pathology depends on the intensity, duration, and the susceptibility of the host. In light infections, the liver appears grossly normal. In heavy infections, a localized dilation of the thickened peripheral bile ducts can be seen on the surface beneath the fibrotic capsule of the liver. Histopathological changes in opisthorchiasis is similar to clonorchiasis previously described.31,32 In the early stage of infection, the biliary epithelium frequently becomes edematous and desquamation may be seen in the areas of tissue in close proximity to the flukes. Periductal infiltrates of mononuclear cells are frequently found, however, inflammation of the bile duct walls is generally only slight in uncomplicated cases. Metaplasia of the biliary epithelial cells transforms into mucin-producing cells (goblet cells) during early infection, and these cells may proliferate to produce many small gland-like structures in the mucosa (adenomatous hyperplasia), leading to a persistent and excessively high mucus content in the bile. Chronic and persistent infections result in a gradual increase in the amount of fibrous tissues, giving the appearance of cholangiofibrosis. As this fibrosis proceeds, the epithelial proliferation becomes milder. These histopathological changes are distinctive features particularly in clonorchiasis, therefore, when proliferation of the ductal epithelium with metaplastic cells and periductal fibrosis are observed in patients in an area of endemic infection, they are highly suggestive of liver fluke infection on histological grounds.32 During the acute or initial stage of infections (<4 weeks post infection), no epithelial hyperplasia or fibrous proliferation has been reported. In chronic infections, however, proliferation of epithelial cells is apparent with the formation of glandular acini, similar to the adenomatous changes in clonorchiasis, and there are varying degrees of periductal fibrosis. Major microscopic changes are confined to the large and medium-sized bile ducts where the flukes are found. The gross and microscopic characteristics of human opisthorchiasis are well established within 7–15 years after O.viverrini infection. In chronic and heavy infections, various degrees of cellular infiltration are caused by superimposed bacterial infection. This may result in suppurative cholangitis, and infection may extend into parenchyma of the liver tissue, causing cholangiohepatitis with abscess formation. In heavy infections with O. viverrini, adult parasites are distributed in the gall bladder, the common bile duct and the pancreatic duct. The worms in large and medium-sized bile ducts induce chronic cholecystitis and when there is superimposed

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bacterial infection, empyema of the gall bladder may result. Unlike clonorchiaisis, stone formation in the bile duct or gall bladder have been rarely observed in a series of post mortem studies.33–35 Parasite-induced mucin-secreting cells produce bile with a high mucin content, which, combined with adult flukes and eggs, serves as a nidus for bacterial superinfection and intrahepatic stone formation.36,37 The mucin-rich bile and the presence of worms and eggs in the bile duct cause cholestasis and provide a favorable environment for secondary bacterial infection, which are of enteric origin, with Escherichia coli being identified most frequently as a pathogen. Enlargement of the gall bladder is commonly found in opisthorchiasis both at autopsy34 and during ultrasonographic studies in patients with O.viverrini infection.36–39

60.1.5 Host Immune Response Although O. viverrini locates in a biliary system and does not invade into the host’s tissue, the infection elicits both local and systemic immune responses. At present, there is little information about specific immune responses to infection, especially cell-mediated immunity and the roles of T-cells and cytokines in protective immunity and the pathogenesis of opisthorchiasis.40 There is marked humoral immune response (IgG, IgA, IgE) to parasite-antigens in the serum and bile of humans and animals infected by the liver fluke.41–44 The level of IgG against crude somatic antigen correlate with the hepatobiliary abnormalities diagnosed by ultrasonography (US), but there is little correlation with the intensity of infection.45 In infected individuals, the level of serum antibodies (IgG, IgA, IgM) change slowly and remains elevated up to 12 months after praziquantel treatment.46 It is not clear whether this reflects long lasting immunological memory or presence of residual parasite antigen derived from treatment or crossantigenic stimulations. Liver fluke DNA has been demonstrated in liver tissue of a CCA patient suggesting that there may be residual parasite material remaining within the liver.47 In the hamster model, antibody responses were first detected as early as 14 days after infection and increased rapidly to a plateau at around 2 months post infection and were relatively stable thereafter.48 Reinfection with O. viverrini metacercariae elicits higher levels of IgG and more severe liver cell damage which correlates with the degree of periductal fibrosis.49 In the case of cellular responses, in vitro proliferative responses of mononuclear cells to mitogen stimulation are unchanged after praziquantel treatment.40 The role of cellular response in biliary pathology in O. viverrini infection is evident since the severity of bile duct inflammation diminished in T-cell deprived hamsters.50 A reduction of an in vitro lymphoproliferative response has been observed in hamsters infected with O. viverrini suggests a degree of immunosuppression.51 In the acute phase of infection in an animal model, evidence of cellular immune response during acute phase was seen as abundant inflammatory cells at 3 weeks post infection with O. viverrini and a marked infiltration of cells included eosinophils, mononuclear cells and neutrophils.48,52 In the chronic phase, from 1 to 6 months of infection, there was an

Molecular Detection of Foodborne Pathogens

increase in mononuclear cells and a decline of eosinophils. Following reinfection, a smaller number of infiltrated inflammatory cells have been observed but mainly in the lymphoid follicles in association with the progression of periductal fibrosis.49 Although evidence presented above clearly showed that infection by O. viverrini stimulated both systemic humoral and cell-mediated immune responses during the course of infection, the significance of these immune responses on protective immunity is not convincing. The fact that some individuals from the endemic area may harbor several thousand worms,53,54 suggests that reinfection does occur and that immune responses fail to prevent reinfection by the same parasite. This conclusion was confirmed when no significant reduction in worm burden was seen in hamsters receiving immune spleen cells and serum compared to the control groups despite a substantial reduction in the fecal egg count.55 It was found that primary infection of hamsters with O. viverrini, whether or not eliminated with praziquantel before challenge, failed to confer any protective immunity to reinfection.56 However, both groups of investigators demonstrated that the previous exposure to O. viverrini depressed egg output in subsequent infections. Egg reduction could be due to biliary obstruction and not necessarily due to immunological damage of the reproductive system of the flukes. The lack of protective immunity is perhaps due to immunosuppression of both cellular and humoral immune responses by O. viverrini infection to unrelated antigens as shown by reduced phytohemagglutinin-induced lymphoproliferation and response to sheep red blood cell stimulation.57 Moreover, when specific antibody was measured in infected animals, the titer was found to be depressed during the late stage of infection, particularly in the heavily infected group.58 The parasites may be resistant to immune damage or able to evade the host defense system. The inability to kill the worms by exposure to serum from infected animals or from patients with opisthorchiasis and the resistance to immune damage may be related to tegumental shedding and repair of the parasite.59 Studies on cellular responses in an animal model indicated that IL-12 which is Th1 type dominates during acute inflammation of bile ducts and is suppressed in chronic infection.60 This study also suggested that TGF-b may play a role in switching from Th1- to Th2-type response (IL-4) in chronic infection stage seen as suppression of cellular immune response. These results confirmed the roles of host response in pathogenesis of opisthorchiasis and CCA.

60.1.6 Clinical Manifestation and Morbidity The recent application of radiology imaging has aided in the diagnosis of subclinical opisthorchiasis as well as associated complications. US appears to be most practical choice for investigation as it is noninvasive and can operate under field conditions.61,62 Characteristic sonographic findings indicated active liver fluke infection and abnormalities, i.e., increased periductal echogenicity, floating echogenic foci in

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gall bladder or gall bladder sludge, diffuse dilatation of intrahepatic bile duct and gall bladder enlargement. Occasionally, biliary stones such as gall bladder stones may be present. The severity of clinical manifestations in opisthorchiasis is correlated with the duration and intensity of infection. According to Sadun,63 severity of infections were classified by intensity of infection (fecal egg count) consisting of light or mild (egg/g feces, epg<1000), moderate or medium (epg<10,000), heavy (epg<30,000) and very heavy infection (epg>30,000) and from autopsy study in northeast Thailand, equivalent worm loads at each intensity classes of <62, <248, <479 and >479 worms, respectively.13 Fecal egg output was used as a parameter to measure worm burden in the biliary system which is associated with severity of hepatobiliary abnormality. Following infection with O. viverrini, the clinical manifestations in the subjects were divided into two groups. Group one was asymptomatic individuals which are the majority of infected people. A continued and repeated infection with no treatment eventually leads to a symptomatic group. But these symptomatic subjects are usually less than 10% of the infected people. Abdominal related symptoms may start to develop particularly in middle age individuals (30–40 years onward) as a consequence of long term chronic infection. The symptoms are considered as mild infection when presenting with abdominal pain, flatulence, right quadrant abdominal pain associated with a burning sensation. In the moderate group the symptoms are more pronounced with liver and gall bladder enlargements but liver function tests have been shown to be normal. In the severe manifestation group, occasional jaundice with fever as a result of biliary obstruction and ascending cholangitis are frequently listed as common clinical manifestations. Liver function tests showed signs of abnormalities such as elevated bilirubin in jaundice cases and sometime liver enzymes may also elevate. In general, observed morbidity in opisthorchiais is usually low, for example, Pungpuk et al.64 reported that only 88 cases (<1%) showed clinical manifestations among 15243 infected people who presented at a hospital in Bangkok. Two large community-based studies by Upatham et al.65,66 reported significant increased frequencies of abdominal pain in the upper right quadrant, flatulence or dyspepsia and weakness associated with increasing intensity of infection. It was estimated that 5–10% of the population had symptoms attributable to infection. Further studies using US have demonstrated strong relationships between gall bladder enlargement, wall irregularities, sludge, enhanced echogenicity of portal vein radicles, and the intensity of infection particularly in moderate to heavy infection.38,67 In another large scale study using a sample size of 12311 individuals, the results confirmed the above findings and showed the relationships in more quantitative terms.53 With the intensity of infection of greater than 3000 epg, the adjusted odd ratio for distended gall bladder and portal vein radicles was between 14–20 and 10–28, respectively.68 Reversibility of these abnormalities as seen by gall bladder size and its regain contractility was observed 11 months after praziquantel treatment.38,69 In

the current situation when infected people were treated with praziquanel, the pattern of hepatobiliary diseases as determined by US has changed considerably. Currently, the heavy infection group (epg>3000) was rare while most of infected people had light infection with epg<1000 and the disease frequency was no longer correlated with intensity of infection as in the past. The ultimate complication in opisthorchiasis is CCA which may constitute within symptomatic subjects but confirmation by radiological investigation is required. It is conceivable that, reduction in prevalence and intensity of O. viverrini as a result of the past and ongoing control program should contribute to lesser morbidity and risk of CCA.

60.1.7 Liver Fluke Induced Cholangiocarcinoma (CCA) Infection and chronic inflammation contribute to carcinogenesis through an inflammation-related mechanism. Infection with hepatitis C virus, Helicobacter pylori and the liver flukes O. viverrini and C. sinensis, are important risk factors for hepatocellular carcinoma, gastric cancer, and CCA, respectively.70 O. viverrini has been classified as a class 1 carcinogen and C. sinensis as a class 2A carcinogen to humans. But, the evidence for O. felineus is insufficient to assess its role in carcinogenesis. CCA, also known as peripheral CCA, is a malignant tumor arising from the biliary epithelium of the intrahepatic biliary tree. It accounts for 10–15% of primary liver cancer and its geographical distribution overlaps with endemic areas of O. viverrini.71 Recent hypotheses on its etiology center on chronic inflammation of the bile ducts and conditions associated with bile stasis are predisposing factors to the development of CCA which include primary sclerosing cholangitis, liver fluke infection and recurrent cholangitis with hepatolithiasis. In Thailand, CCA is 12 times more common in the northeast, the endemic area of O. viverrini, than in regions with low prevalence of O. viverrini.72 The age-standardized incidence of CCA in Khon Kaen of 84.6 and 36.8 per 100000 for male and female, respectively, was the highest in the world.71 Cancer epidemiological studies revealed that evidence of current or past experience of liver fluke infections, in terms of intensity of infection, high serum antibody against the liver fluke, are risk factor of CCA.73,74 The frequency of suspected CCA is positively associated with fecal egg count where the odd ratio14.1 for CCA was found in a group with epg>6000, which is equivalent to the presence of >120 worms in the biliary tract.53 Additional risk factors reported in CCA were areas of residence, alcohol consumption, age, and sex and recently host genetic polymorphism of glutathione S-transferase enzyme (GSTM1) in association with seropositivity for opisthorchiasis, suggested that host genetic factors also contribute as risk factors of CCA. These findings suggest a classic gene-environment interaction which plays crucial roles in individual susceptibility to cancer. Histological features of CCA in the endemic area of liver fluke infection is similar to that described in nonendemic regions.36 Tumor cells originated from surface epithelial cell

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lining the peripheral bile duct invade the sinusoids of the adjacent liver parenchyma to form a tumor nodule. In most clinical cases, the disease is in advanced stage and usually appears as a large, single, white firm tumor with a distinct margin in the non-cirrhotic liver.75,76 The most common histopathology of CCA (>95%) is adenocarcinoma showing a glandular and/or papillary structure with a variable fibrous stroma. There is no dominant histological type of CCA in both endemic and non-endemic areas of the liver fluke.77 A typical hyperplasia and dysplasia that lie adjacent to CCA are likely to be precancerous lesions.78 Most authors of the earlier studies suggest that liver flukes mediate tissue damage directly by mechanical and chemical irritation, but some studies postulated that parasite-specific immune responses may play a major role.31,50,55,79 At present, although the underlying enhancement of neoplasia by liver flukes is not fully understood, carcinogenesis is a multi-stage process in which many factors are likely to be involved. However, the mechanistic explanation is gradually becoming clearer particularly the hypothesis that the chronic infection and inflammation is a risk factor of carcinogenesis due to the possible role of nitric oxide.80 During liver fluke infection, macrophages, and other cell types (e.g., mast cells, eosinophils, epithelial cells), activated by parasite-specific T-cell and cytokines, synthesize NO from L-arginine via the induction of nitric oxide synthase (iNOS) in order to kill the parasite. NO is not only cytotoxic, but also genotoxic at physiological concentrations. Excess NO production plays a crucial role in a variety of pathological processes, including cancer.80 NO reacts with superoxide (O2•–) to form peroxynitrite (ONOO–), a highly reactive species causing nitrative and oxidative DNA damage. ONOO– can mediate the formation of 8-oxodG and 8-nitroguanine, a marker of nitrative DNA damage.81–83 Endogenous nitrosation caused by liver fluke infection has been studied in both animal models and humans. In O. viverrini infected people, there were significant increases in plasma and urinary nitrate and N-nitrosoproline (NPRO) in urine after proline loading, nitrate, and NPRO, the indicators of endogenous NO synthesis and nitrosation reactions respectively.84,85 The site of the endogenous reaction was proven to be extragastric, i.e., within the inflamed bile duct.86 Direct measurement of the carcinogenic product of nitrosation reaction, N-nitrosodimethylamine (NDMA), excreted in the urine, appeared to be associated with in vitro lymphoproliferative responses to liver fluke antigens; after the flukes were removed following praziquantel treatment, the association disappeared.87 In multivariate analyses, NDMA levels were related to urinary nitrates, stimulation indices for two T-cell responses to two parasite antigens (MW 37 kDa and 110 kDa) and gall bladder dimensions. In active infection with O. viverrini, NDMA is further activated by some isoforms of cytochrome P450 (CYP) enzymes, mainly CYP2E1 and CYP2A6 to become ultimately carcinogenic as shown in human and animal models.88,89 This increase in CYP 2A6related enzyme activity may represent an important mechanistic link between inflammatory products of chronic liver

Molecular Detection of Foodborne Pathogens

fluke infection and the high risk of CCA faced by infected individuals. A recent study in humans showed that 8-oxodG levels in O. viverrini-infected subjects were higher than in healthy subjects and this DNA adduct significantly decreased 2 months post praziquantel treatment and were comparable with levels in healthy subjects 1 year after treatment.90 Therefore, NO, and ROS may mediate 8-oxodG and 8-nitroguanine formation triggered by O. viverrini infections through chronic inflammation reactions. O. viverrini causes the formation of oxidative and nitrative DNA adducts as proven through the Toll like receptor 2 (TLR2)-mediated pathway leading to the activation of nuclear factor κB (NF-κB), which is a key player in tumor promotion in inflammation associated cancer.91 Additionally, proliferating cell nuclear antigen (PCNA), a cofactor for DNA polymerase, is associated with DNA replication and long-patch base excision repair, accumulated in the epithelium of bile ducts after repeated O. viverrini infection, supporting the hypothesis that cell proliferation was promoted by inflammation mediated DNA damage.92 It was shown that the responses of a fibroblast cell line, NIH-3T3, to excretory/ secretory (ES) product(s) released from O. viverrini in a noncontact coculture condition, had a marked increase in cell proliferation.93 Treatment with praziquantel not only reduced bile duct inflammation via TLR-2 mediated pathway but also oxidative and nitrative DNA damage when assessed during 1–3 weeks post treatment and thus may help prevent carcinogenesis.94 However, subsequent studies suggest that short-term praziquantel treatment induces inflammation and resulting oxidative and nitrative stress through O. viverrini antigen release.95 Praziquantel treatment clearly helps to inhibit iNOS-dependent DNA damage through the elimination of the parasites, but also act as a potential anti-inflammatory effect. A possible mechanism of liver fluke-associated CCA development has been proposed and it contains four stages similar to those in colon cancer.89,96,97 Firstly, exposure to risk factors (opisthorchiasis) leads to chronic inflammation and/or cholestasis through the biochemical and/or mechanical processes. Secondly, genotoxic events may be the consequence of NO overproduction by inflammatory cells. NO and oxygen radicals can inactivate and injure the lipid, protein, DNA, and carbohydrate within the cells, via direct oxidative damage. Free radicals caused by inflammatory cytokines can induce many subsequent events, such as the activation of gene expression, alteration of detoxification gene expression and activation of carcinogen metabolism to form ultimate carcinogens, which enhance DNA damage. Moreover, overproduction of NO has been reported to down regulate lymphocyte proliferation. Thirdly, dysregulation of DNA repair and apoptosis as well as genotoxic events with DNA damage usually lead to either DNA mismatch repair or if the damage is not repair, cell death will follow through apoptosis. Finally, histological changes to CCA may occur in the manner of “hyperplasia-dysplasia-carcinoma” model which has been proposed for colon and gastric cancers.97

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60.1.8 Epidemiology, Prevention, and Control After the discovery of O. viverrini in 1915 in Chiang Mai Thailand, subsequent studies in 1955 by Sadun63 confirmed the identity of the liver fluke that was found to be endemic in Thailand was in fact O. viverrini. The pattern of infection in populations in endemic communities often showed that the prevalence and intensity increased with age of the population thus middle aged people tend to have high prevalence and intensity of infection. Likewise associated morbidity also frequently occurred in this age group. The distribution of worm burden in humans is not uniform, since the majority of parasites are harbored by a minority group of people. In addition, there is evidence of predisposition to heavy infection in which heavily infected individuals have a tendency to return to their pretreatment infection levels after treatment.98 These epidemiological characteristics have important bearing on the subsequent design and planning of control programs. While snail control to disrupt the life cycle is not an attractive option for control of opisthorchiasis, chemotherapy by using praziquantel is the most common means for large scale parasite control as well as individual treatment. The treatment program varies according to the local situation and constraint in each country. In Thailand for example, a control program was implemented via selective treatment approach (treatment given to infected individual as opposed to mass treatment where treatment given is to all members in an endemic community). Although the efficacy of praziquantel is relatively high (90–95%) and to date no evidence of drug resistance has been reported, reinfections are a common phenomenon in many endemic areas.98,99 Therefore parasite control by chemotherapy alone may not be successful and a combined measure of treatment, sanitary improvement and health education, is recommended.100 A self-reliance control program was adopted at the beginning of the campaign during 1980–1987 in Thailand. This approach has contributed to significant reductions in the prevalence and intensity of infection.9 In addition to chemotherapy, health education to avoid eating raw or under cooked fish preparation is essential to reduce and prevent infection. This eating behavior which is deep rooted, a part of culture and tradition for decades in endemic areas of O. viverrini i.e., northeast Thailand and Lao PDR, is not easy to change. Thus the health education message should be culturally sensitive for a given community. For the current population of adults who have been exposed to infection and are at risk of having hepatobiliary disease and CCA, health education may need to be focused on the importance of reinfection which can further increase the risk of CCA. Another approach for control is targeting the young generation such as school children, hence realize the future goal of a parasite free generation having low or no risk of CCA. Because of the growing fishery export trade from endemic areas of foodborne trematode to nonendemic areas, both WHO and FAO are aware of the importance of an integrated, multidisciplinary approach to food safety and quality, which considers the entire food chain. In the case of aquaculture,

the food chain approach required is a responsibility that the fish supplied is safe for human consumption by all involved with the production, processing, trade, and consumption of fish. Development and implementation of Good Aquaculture Practice (GAP), Good Hygienic Practice (GHP) and Hazard Analysis Critical Control Point (HACCP) are recommended for fish farming in order to reduce metacercarial contamination of fish and clean fishery products.4 These measures are possible in intensive, industrial aquaculture practices but for small scale subsistence fish farming a balance between the operation cost and potential profit should be considered. In addition, preventive measures can be achieved by several techniques to reduce the risk of O. viverrini in fish. These included freezing (e.g., at –20oC for 7 days) or heat treatment (e.g., cooking).

60.1.9 Laboratory Diagnosis The ideal gold standard diagnosis in medical and veterinary helminthiasis is a demonstration of life stages of the parasite in the given hosts. In some circumstances when recovery or observation of adult stages is not feasible, ova or larval stages are indirect evidence for the presence of infection and this is also accepted as a gold standard diagnosis in parasitology. In opisthorchiasis, detection of eggs by fecal examination is considered as a standard diagnosis in man and animal hosts. Herein only commonly used methods for diagnosis of opisthorchiasis are reviewed with emphasis on their advantages and limitations. Improved diagnostic methods for the current transmission situation are required for better parasite control and therefore to enable us to reduce disease burden. 60.1.9.1  Parasitological Techniques Demonstrations of eggs in feces, bile, duodenal fluid or recovery of adult flukes during transhepatic stent implantation or directly from liver in post mortem autopsy are the gold standard diagnosis for O. viverrini infection. Fecal examination is the most common method used for routine diagnosis. The sensitivity of the diagnosis depends on method of fecal examination and also the experience of the microscopist. The techniques frequently employed for diagnoses of liver fluke infection are the formalin-ether concentration technique, the Kato thick smear and Stoll’s dilution egg count technique. Other standard techniques such as a simple direct smear or simple sedimentation techniques are also used in certain circumstances depending on the purpose of the study. Single examination by these techniques seems to have a high falsenegative rate, especially with light infections or those with a history of recent treatment. Repeated stool examination is recommended to improve the sensitivity of the examination. Although repeated examination increases sensitivity, repeated or multiple stool sample collections is costly and requires more compliance which may not be feasible in the field. The formalin-ether concentration technique is considered to be relatively sensitive compared to other techniques, since fecal material and fat are removed; the resulting sediment

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is clear and the eggs are easily seen microscopically. This technique can be modified to be quantitative if the weight or volume of the stool sample is known and the fecal egg output calculated. The other advantage is that the stool samples, once fixed in formalin, can be kept for several months prior to processing and examination. The typical amount of stool sample taken for processing by this technique is 1–2 g which is 1000–2000 more than the direct simple smear technique. In light infection cases with low worm burdens, egg detection in the stool is unreliable. The sensitivity of fecal examination by formalin-ethyl acetate concentration and Stoll’s egg count technique are dependent on worm burden in the liver. Based on recovery of adult O. viverrini directly from liver at autopsy (n=139), prevalence determined by worm recovery was found to be 81.29% This was significantly greater than either formalin-ether concentration (61.87%) and Stoll’s dilution egg count (66.91%) techniques.13 A detailed analysis revealed that in cases of worm burden of less than ten worms per liver, as much as 68–72% were egg negative in stools. At higher worm loads of 10–19 worms, the proportion of false negatives became 23–29%, while at a higher worm burden >20, the rate of egg detection was comparable with worm recovery method. The Kato thick smear technique is commonly used for routine diagnosis of opisthorchiasis in Thailand and Laos, and it is also widely used for clonorchiasis in Korea and China. It is believed to be a sensitive and reliable technique comparable to the formalin-ether concentration technique and more sensitive than the simple sedimentation technique particularly for clonorchiasis.101 When the amount of stool specimen is measured or weighed it is then known as the Kato-Katz technique (KK). However, the identification of eggs in the Kato thick smear method requires experience to distinguish eggs of O. viverrini from eggs of intestinal trematodes. The other common fecal examination method is the Stoll’s dilution egg count technique in which sodium hydroxide is used to prepare the fecal suspension. Sensitivity has been reported to be slightly inferior to KK technique, however, it may be suitable for assessment of the intensity of O. viverrini.102 Since there are several species of foodborne trematodes belonging to the Opisthorchiidae, Heterohyidae, and Lecithodendriidae families which have similar egg morphology, recognition of egg morphology is essential for correct identification. In Thailand and Laos the latter two families are collectively referred to as minute intestinal flukes (MIF) because of their small size compared to O. viverrini.14 The Lecithodendriidae contains Phaneropsolus bonnei and Prostodendrium molenkampi while the Heterohyidae contains Haplorchis taichui, H. pumilio and Stellanchasmus falcatus.103,104 Within the heterophyid, Metagonimus spp, Heterophyid spp, Haplorchis spp and Pygidiopsis spp were found in Laos and Korea.1 The adult worms of liver flukes namely O. viverrini, C. sinensis and O. felineus are distinguishable but their eggs are similar morphologically. Moreover, eggs of heterophyids are very similar in all the three liver flukes; thus, adult worms are required for identification.5

Molecular Detection of Foodborne Pathogens

Under light microscopy, eggs of O. viverrini are characterized by rough and thick egg shells; and by scanning electron microscopy the egg shell shows a mask melon pattern. This can be differentiated from P. bonnei and P. molenkampi which have smooth and thin egg shells.14 In areas where O. viverrini coexist with Heterophyid fluke i.e., Haplorchis spp, a term “O. viverrini-like egg” is sometime used.105 This dilemma clearly reflects the limitation of the diagnostic approach based solely on morphological criterion and there is a great need for alternative diagnosis with high sensitivity as well as specificity. 60.1.9.2  Immunological Techniques Serodiagnosis, by detecting anti-parasite antibodies, plays roles in diagnosis of several blood and tissue parasites and is widely used to compliment stool examination. In the past decade, serological tests have been developed so that the sensitivities and specificities of the tests have greatly improved. Currently, there are several techniques available such as an intradermal test, immunoelectrophoresis (IEP), indirect hemagglutination assay (IHA), indirect fluorescent antibody test (IFAT) and indirect enzyme-linked immunosorbent assay (indirect ELISA).40 Among these tests, ELISA is the most favored method, but its sensitivity and specificity vary depending on the nature of the antigen used in the detection system. Different parasite components are prepared and utilized as antigens for instance, adult somatic extracts, excretory, and secretory (ES) substance, and surface tegument and egg antigens. In opisthorchiasis, adult somatic extracts have been used for IEP and ELISA with a sensitivity of 76–100% for IgG and lower sensitivities for IgA and IgE.106,107 The 89 kDa protein, most prominent in the ES antigen of O. viverrini, was claimed to be specific with immunodiagnostic potential; however, it also cross reacted with C. sinensis.108 The affinity-purified egg antigen using specific monoclonal antibody, to detect serum antibody gives a high sensitivity and specificity.107,109 Using liver fluke-derived crude or purified antigen in this test platform has been less satisfactory because of cross reactive antigens. Attempts to produce recombinant antigen for IgG antibody detection by ELISA, such as the recombinant O. viverrini egg shall protein (rOVESP), was found to be more sensitive and more specific than native ES antigen.110 However, irrespective of the quality of antigens, one drawback of these antibody detection methods is the inability to differentiate past and current infection status since antibodies in opisthorchiasis as well as clonorchiasis persist in the infected hosts for months or years even after curative treatment.46,111 More recombinant antigens are becoming available, therefore, there are possibilities that the serological tests could be improved. Recent molecular genetic studies of O. viverrini have begun to disclose a range of genes encoding various types of proteins, some of which may be potential candidates for recombinant antigens for serodiagnosis as well as for studying their biological function in the worm.112 Unlike antibody detection, antigen detection is informative in relation to the current state of infection. For this

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approach, several monoclonal antibodies to O. viverrini antigens were produced against different antigenic proteins such as 16 kDa tegumental protein, 89 kDa glycoprotein, and 90 kDa somatic protein and these monoclonal antibodies were used for coproantigen detection with varying success with a sensitivity of 31–57% and a specificity of 70–100%.113,114 By employing the monoclonal antibody specific to 89 kDa antigen of O. viverrini in the monoclonal antibody based-ELISA (Mab-ELISA) for antigen detection in stool specimen, diagnosis by the Mab-ELISA was found to be more sensitive than the conventional fecal examination particularly light infection group (epg<500).113 This finding is in concordance with results from an autopsy study which demonstrated that fecal eggs were not detectable in a group with <20 worms in the liver.13 It may be possible that Mab is sensitive enough to detect secretory product from a few adult worms when eggs in the feces are not detected. Although this is a promising approach to pursue and develop further into a simple diagnostic kit, a supply of desirable Mab is no longer available. Recently new clones of Mab against ES product of O. viverrini, recognizing the molecular weight of 45–110 kD were established and were employed to develop the Mab-ELISA for fecal antigen. This latest Mab-ELISA gave the sensitivity of 69.1% and the specificity of 39.1% (unpublished). Moreover, similar to Sirisinha et al.113 a considerable number of cases have been discovered by the present antigen detection in egg-negative subjects (unpublished). Verification of these positive tests by antigen detection is now underway by comparing these results with those of DNA-based diagnostic methods. This analysis may shed light on the development of alternative diagnostic methods in addition to classical fecal examination. 60.1.9.3  Molecular Techniques Parasitological and immunological diagnostic methods have problems in terms of their specificities but molecular methods are particularly advantageous, specifically on specificity. In opisthorchiasis, a specific DNA probe designed from repeated DNA elements (satellite DNA) of 334 bp has been used for the detection of egg DNA.115 Subsequently, a PCRbased detection of O. viverrini in human stools, based on a pair of primers complementary to the same target DNA of O. viverrini, was used to detect egg DNA.116,117 From these studies, a specificity of 97.8% and a sensitivity of 100% was achieved in moderate to severe infections (epg>1000) but in light infections (epg<200), the sensitivity was reduced to 68.2%. Application of this PCR-based method for the detection of O. viverrini DNA in stool samples from Laos yielded sensitivity of approximately 50% in samples with egg count >1000 epg.118 The authors suggested that the presence of PCR inhibitor in the fecal specimen may influence the sensitivity of the diagnosis rather than the diagnostic reference related issue. Recent improvement of fecal DNA preparation using cetyltrimethylammonium bromide (CTAB) to remove PCR inhibitor was employed and when compared with commercial stool kit method, the sensitivity of the new test was 79.3% compared with 44.8% in the previous

method.119 In this study PCR-positive tests were identified in several parasite negative cases by parasitological method (28.6%), indicating its advantage in diagnosis of individuals with light infection. A new O. viverrini-specific primer pair was recently established for the PCR test and the sensitivity of the test was 10–12 ng of adult worm DNA and three metacercariae in fish sample.120 In addition, species-specific PCR tests to identify the species of liver fluke are now available for O. viverrini,117,121 O. viverrini, C. sinensis,122 and O. felineus.123 It is worth noting that as these molecular diagnostic tests become extremely specific they will play significant roles in the accurate assessment of cures as well as the rate of reinfection of O. viverrini in endemic areas. It is possible to identify the parasite species from various life stages of the parasite including eggs, metacercaria as well as adult worms. Owing to their sensitivity and specificity, DNA based methods are being recognized as potential tools to detect different life stages of parasites especially metacercaria in fish. With aquaculture of fish playing a vital role in international trade, assurance of the safety of fish for human consumption has assumed significance.11 Application of HACCP can be greatly enhanced by the application of robust DNA-based techniques to assess the effectiveness of these measures to reduce risks at various stages of food production. PCR has the advantage of being able to screen large number of samples in a relatively short time with high sensitivity and specificity. Conventional techniques to detect metacercariae in fish require enzyme digestion of the tissue, sedimentation of metacercariae and microscopy. In this context, the PCR assay developed by Pavarthi et al.120 offers a reliable method for the detection of O. viverrini metacercariae in fish for food safety purposes and human consumption.

60.2  Methods Several molecular detection methods for human opisthorchiasis have been developed over the past decades. We describe below methods of detection of O. viverrini DNA in stool samples using PCR technology with details on preparation of DNA template from stool samples and PCR amplification conditions.

60.2.1 Preparation of O. viverrini DNA using a Commercial Kit DNA is prepared from stool sample using QIAamp® DNA Stool Mini Kit (Qiagen, Germany).118 Reagents: Normal saline (NS) (0.85%NaCl), ethyl acetate, 0.5 N NaOH, 3 M sodium acetate, QIAamp® DNA Stool Mini Kit (Qiagen, Hilden, Germany), and proteinase K. Procedure: (1) Mix fecal samples (100–500 mg) with 1 ml of NS and 200 µl of ethyl acetate. (2) Centrifuge the mixture at 8000 rpm for 5 min.

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(3) Wash the pellet with 1.5 ml of NS, mixed with a vortex and incubated in 500 µl of 0.5 N NaOH for 60 min at room temperature. (4) Bake the mixture in autoclave oven at 121oC for 60 min. (5) Vortex the mixture and centrifuge at 8000 rpm for 5 min. (6) Examine samples for the presence of broken eggs after autoclavation using microscope. (If broken egg shells are not seen, repeat step 4 again.) (7) Transfer the supernatant to a 1.5 ml centrifuge tube. (8) Add 250 µl of 3 M sodium acetate and adjust pH to 7–8. (9) Add half a tablet of one InhibitEX (QIAamp® DNA Stool Mini Kit; Qiagen, Hilden, Germany) to reduce potential inhibition and centrifuge the mixture for 5 min at 14000 rpm. (10) Transfer the supernatant into a new tube, and discard the pellet. (11) Add 10 µl of proteinase K (20 mg/ml) to the supernatant. (12) After adding 200 µl Buffer AL supplement in the QIAamp® DNA Stool Mini Kit and mixing, incubate the suspension at 70oC for 10 min. (13) Add 200 µl of absolute ethanol, and clean the DNA solution and elude using a QIAamp® DNA Stool Mini Kit (Qiagen, Hilden, Germany) according to the instructions of the manufacturers. (14) Use 5 µl of the extracted DNA for PCR.

60.2.2 Preparation of O. viverrini DNA using a CTAB Method O. viverrini DNA can be also prepared from stool samples using CTAB.119,120 The purpose of this method is to isolate DNA from O. viverrini egg in human stool samples. This manual protocol is modified from the DNA extraction method described by Ausubel.124 Reagents: Normal saline (NS) (0.85% NaCl), ethyl acetate, 0.5 N NaOH, 3 M sodium acetate, 10× TE (pH 8.0), proteinase K, 10% SDS, CTAB, chloroform:isoamyl alcohol (24:1), and phenol:chloroform:isoamyl alcohol (25:24:1) Procedure: (1) Mix fecal samples (approximately 500 mg) with 4 ml of 0.85% NaCl and 400 μl of ethyl acetate in 15 ml screwed cap tube. (2) Vortex the sample vigorously and centrifuge at 4000 rpm for 10 min. (3) Resuspend the pellets in 1 ml of 0.5 N NaOH and incubate for 60 min at room temperature. (4) Autoclave the mixture at 121oC for 60 min. (5) Vortex the sample vigorously and centrifuge at 4000 rpm for 10 min.

Molecular Detection of Foodborne Pathogens

(6) Evaluate the sample by microscopy for the presence of broken egg shell prior to the next step. (If broken egg shells are not seen, repeat step 4 again.) (7) Transfer the supernatant to a 1.5-ml microcentrifuge tube, add 56 μl of 1× TE (pH 8.0) and adjust to pH 7–8. (8) Add 30 μl of 10% SDS and 3 μl of proteinase K (20 mg/ml) to the supernatant and incubate at 65oC for 1–2 h. (9) Add 100 μl of 5 M NaCl and 80 μl of CTAB to the supernatant and incubate at 65˚C for 30 min (0.7 M NaCl and 0.1 volume of CTAB). (10) Extract the mixture twice with an equal volume of chloroform:isoamyl alcohol (24:1) and once with phenol:chloroform:isoamyl alcohol (25:24:1). (11) Precipitate DNA from aqueous phase with 0.6 volume of isopropanol. (12) Pellet the DNA by centrifugation, wash once with 70% ethanol, air dry and dissolve in 1× TE buffer. (13) Use 2 μl of the extracted DNA for PCR.

60.2.3 PCR Detection of O. Viverrini from Stool Samples Caution: It is important to have separate areas in the PCR lab., for sample preparation, pre-PCR setup and post-PCR detection, with an exclusive set of automatic pipettes for each area. All PCR reagents are aliquoted into individual tubes and the master mix is prepared to reduce contamination. The reproducibility of the method is checked by amplification of the positive control (pooled O. viverrini genomic DNA) and negative control in parallel with other samples. 60.2.3.1 Use of pOV-6 Primers The pOV-6 primers were originally designed for the detection of O. viverrini in experimental infected hamsters.117,125 Application in human opisthorchiasis, has shown the assay to have a specificity of 97.8%, a sensitivity of 100% in moderate-severe infections (more than 1000 epg feces) and a maximum sensitivity of 68.2% in the detection of light infections (epg<1000) with a detection limit of 200 eggs.116 Reagents and equipment: DNA thermal cycler (MJ PTC-2000), PCR kit (Ready-To-GoTM PCR Beads, Amersham Pharmacia Biotech), primers (pOV-6 forward: 5′-CTGAATCTCTCGTTTGTTCA-3′ and pOV-6 reverse: 5′-GTTCCAGGTGAGTCTCTCTA-3′), 2× master mix (Taq DNA polymerase, 10 mM Tris-HCl, 50 mM KC1, 1.5 mM MgCl2, 200 mM dNTP and stabilizers) Procedure: (1) Prepare PCR mixture (50 µl) containing 25 µl of 2× master mix, 2.5 µl (25 pmol) each of pOV-6 forward and pOV-6 reverse primers, 5 µl of template DNA (QIAamp method), and 15 µl of distilled H2O. (2) Perform PCR amplification with one cycle of 94oC for 5 min; 30 cycles of 94oC for 30 sec, 52oC for 30 sec, 72oC for 45 sec.

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(3) Separate the PCR product by electrophoresis in a 1% agarose gel with ethidium bromide using 1× TBE buffer pH 8.0. (4) Visualize the specific 330 bp PCR products under UV transilluminator. 60.2.3.2 Use of Trem25-F and OV25-4R Primers The Trem25-F and OV25-4R primers were originally developed from a DNA fragment amplified with RAPD-PCR.120 Reagents and equipment: DNA thermal cycler (MJ PTC-2000), primers (Trem25-F: 5′-ACAGGAAGTGGAA CCGTGTC-3′ and OV25-4R: 5′-AATGAACGGAAATC GTGACC-3′), Taq DNA polymerase, dNTP, 10× PCR buffer (20 mM Tris-HCl (pH 8.4), 50 mM KCl and 1.5 mM MgCl2). Procedure: (1) Prepare PCR mixture (30 µl) containing: Template DNA (CTAB method)

  2 µl

dNTP (10 mM)

  0.3 µl

Primer Trem25-F (10 pmol)

  1 µl

Primer OV25-4R (10 pmol)

  1 µl

Tag polymerase (2 units)

  0.5 µl

10× PCR buffer

  3 µl

Distilled H2O

22.2 µl

(2) Perform PCR amplification with the following cycling program: one cycle of 95oC for 5 min; 35 cycles of 94oC for 1 min, 60oC for 1 min, and 72oC for 1 min; one cycle of 72oC for 5 min. (3) Subject the PCR product to agarose electrophoresis using 1.6% agarose gel containing 0.5 µg/ml ethidium bromide in 1× TBE buffer. (4) Visualize the specific 229 bp product with UV transiluminator.

60.3 Conclusions and Future Perspectives O. viverrini is a foodborne trematode, which is an important health problem because of the massive numbers of people infected and its serious morbidities such as hepatobiliary diseases and CCA. Although infections are identified throughout southeast Asia, the epi-center is northeast Thailand, where high prevalence coexists with a high incidence of CCA. Such information in nearby endemic countries, i.e., Laos and Cambodia is currently lacking. O. viverrini requires three hosts to complete its life cycle. Human infection occurs through consumption of the infective metacerariae contained in undercooked freshwater fish. Once eggs reach water, they are ingested by snail intermediate hosts in the genus Bithynia. The parasites multiply asexually in the snail and release the free swimming cercariae into water which subsequently enter the fish intermediate hosts. Common fish intermediate hosts include

several species of cyprinid fish, naturally bred fish species as well as cultured freshwater fish, all of which can act as the source of human infection. Although pathogenesis of opisthorchiasis is complex, recent studies have revealed that the pathogenesis feature in opisthorchiasis is largely a consequence of immunomodulation during the acute and chronic phase of infection. For pathogenesis of CCA, chronic inflammation plays a central role in carcinogenesis through oxidative and nitrative DNA damage which initiates tumorigenesis. While CCA is fatal and curative treatment is not forthcoming, opisthorchiasis is effectively cured by treatment with praziquantel. Praziquantel treatment eliminates not only O. viverrini but also reduces associate morbidities and risk of CCA. A desirable approach for the prevention of infection by O. viverrini is by the consumption of cooked fish. Unfortunately, the campaign, to date, for changes in the eating habits of people appears to have met with little success. Diagnosis of opisthorchiasis for appropriate treatment is therefore essential to prevent morbidity and disease complication. The standard conventional diagnosis is the parasiotological method of demonstrating eggs in feces or bile specimens. Several immunological diagnostic methods by antibody or antigen detection are available but careful interpretation of the test result is required. Molecular diagnostic methods have been introduced as an alternative method for diagnosis with the advantage of high specificity and sensitivity. Diagnosis by detection of parasite DNA in fecal specimens is a promising method since it identifies positive tests even in a considerable number of egg-negative specimens and equally sensitive in egg positive cases. The ability to discover positive cases in apparently egg negative cases as determined by the conventional method, has demonstrated that this PCRbased diagnostic approach is useful for both individual and mass screening diagnosis. It also provides an additional tool for evaluation of the chemotherapy program. With fish aquaculture playing a vital role in international aquaculture trade, this PCR-based method serves as a tool for fish inspection for metacercariae, a significant issue in food safety. The disadvantage of molecular diagnostic methods for detection of the parasite DNA is the requirement of specific molecular instrumentation, thus the development of the detection system into a simplified kit format for routine field and laboratory use is required.

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Opisthorchis 48. Sripa, B., and Kaewkes, S. Localisation of parasite antigens and inflammatory responses in experimental opisthorchiasis. Int. J. Parasitol., 30, 735, 2000. 49. Pinlaor, S. et al. Hepatobiliary changes, antibody response, and alteration of liver enzymes in hamsters re-infected with Opisthorchis viverrini. Exp. Parasitol., 108, 32, 2004. 50. Flavell, D.J., and Flavell, S.U. Opisthorchis viverrini: pathogenesis of infection in immunodeprived hamsters. Parasite Immunol., 8, 455, 1986. 51. Wongratanacheewin, S. et al. Immunodepression in hamsters experimentally infected with Opisthorchis viverrini. J. Helminthol., 61, 151, 1987. 52. Thamavit, W. et al. Effects of dimethylnitrosamine on induction of cholangiocarcinoma in Opisthorchis viverrini-infected Syrian golden hamsters. Cancer Res., 38, 4634, 1978. 53. Haswell-Elkins, M.R. et al. Cross-sectional study of Opisthorchis viverrini infection and cholangiocarcinoma in communities within a high-risk area in northeast Thailand. Int. J. Cancer, 59, 505, 1994. 54. Haswell-Elkins, M.R. et al. Distribution patterns of Opisth­ orchis viverrini within a human community. Parasitology, 103, 97, 1991. 55. Flavell, D.J. et al. Opisthorchis viverrini: partial success in adoptively transferring immunity with spleen cells and serum in the hamster. J. Helminthol., 54, 191, 1980. 56. Sirisinha, S. et al. Attempts to induce protective immunity in hamsters against infection by a liver fluke of man (Opisthorchis viverrini). Parasitology, 86, 127, 1983. 57. Wongratanacheewin, S. et al. Effect of praziquantel treatment on antibody levels and lymphoproliferative responses in patients with opisthorchiasis. Southeast Asian J. Trop. Med. Public Health, 19, 109, 1988. 58. Sirisinha, S. et al. Humoral immune responses in hamsters infected with Opisthorchis viverrini. Southeast Asian J. Trop. Med. Public Health, 14, 243, 1983. 59. Sirisinha, S., Wongratanacheewin, S. Immunization of hamsters against Opisthorchis viverrini infection. Southeast Asian J. Trop. Med. Public Health, 17, 567, 1986. 60. Jittimanee, J. et al. Cytokine expression in hamsters experimentally infected with Opisthorchis viverrini. Parasite Immunol., 29, 159, 2007. 61. Mairiang, E., and Mairiang, P. Clinical manifestation of opisthorchiasis and treatment. Acta Trop., 88, 221, 2003. 62. Choi, M.S. et al. Correlation between sonographic findings and infection intensity in clonorchiasis. Am. J. Trop. Med. Hyg., 73, 1139, 2005. 63. Sadun, C.M.E.H. Studies on the epidemiology of the human intestinal fluke, Fasciolopsis buski (Lankester) in central Thailand. Am. J. Trop. Med. Hyg., 2, 1070, 1953. 64. Pungpak, S. et al. Ultrasonographic study of the biliary ­system in opisthorchiasis patients after treatment with praziquantel. Southeast Asian J. Trop. Med. Public Health, 20, 157, 1989. 65. Upatham, E.S. et al. Relationship between prevalence and intensity of Opisthorchis viverrini infection, and clinical symptoms and signs in a rural community in north-east Thailand. Bull. WHO, 62, 451, 1984. 66. Upatham, E.S. et al. Morbidity in relation to intensity of infection in Opisthorchiasis viverrini: study of a community in Khon Kaen, Thailand. Am. J. Trop. Med. Hyg., 31, 1156, 1982. 67. Dhiensiri, T. et al. Roentgenographically controlled healing of gallbladder lesions in opisthorchiasis after praziquantel treatment. Arzneimittelforschung, 34, 1175, 1984.

825 68. Elkins, D.B. et al. Cross-sectional patterns of hepatobiliary abnormalities and possible precursor conditions of cholangiocarcinoma associated with Opisthorchis viverrini infection in humans. Am. J. Trop. Med. Hyg., 55, 295, 1996. 69. Mairiang, E. et al. Reversal of biliary tract abnormalities associated with Opisthorchis viverrini infection following praziquantel treatment. Trans. R. Soc. Trop. Med. Hyg., 87, 194, 1993. 70. IARC. Infection with liver flukes (Opisthorchis viverrini, Opisthorchis felineus and Clonorchis sinensis). IARC Monogr. Eval. Carcinog. Risks to Hum., 61, 121, 1994. 71. Parkin, D.M. et al. Cholangiocarcinoma: epidemiology, mechanisms of carcinogenesis and prevention. Cancer Epidemiol. Biomarkers Prev., 2, 537, 1993. 72. Srivatanakul, P. et al. Epidemiology of liver cancer: an overview. Asian Pac. J. Cancer Prev., 5, 118, 2004. 73. Lim, M.K. et al. Clonorchis sinensis infection and increasing risk of cholangiocarcinoma in the Republic of Korea. Am. J. Trop. Med. Hyg., 75, 93, 2006. 74. Honjo, S. et al. Genetic and environmental determinants of risk for cholangiocarcinoma via Opisthorchis viverrini in a densely infested area in Nakhon Phanom, northeast Thailand. Int. J. Cancer, 117, 854, 2005. 75. Uttaravichien, T. et al. Intrahepatic cholangiocarcinoma in Thailand. J. Hepatobiliary Pancreat. Surg., 6, 128, 1999. 76. Uttaravichien, T., and Buddhisawasdi, V. Experience of nonjaundiced cholangiocarcinoma. Hepatogastroenterol., 37, 608, 1990. 77. Pairojkul, C. et al. Multistage carcinogenesis of liver-flukeassociated cholangiocarcinoma in Thailand. Princess Takamatsu Symp., 22, 77, 1991. 78. Hou, P.C. The relationship between primary carinoma of the liver and infestation with Clonorchis sinesis. J. Pathol. Bacteriol, 72, 239, 1956. 79. Haswell-Elkins, M.R. et al. Immune responsiveness and parasite-specific antibody levels in human hepatobiliary disease associated with Opisthorchis viverrini infection. Clin. Exp. Immunol., 84, 213, 1991. 80. Ohshima, H., and Bartsch, H. Chronic infections and inflammatory processes as cancer risk factors: possible role of nitric oxide in carcinogenesis. Mutat. Res., 305, 253, 1994. 81. Kawanishi, S. et al. Oxidative and nitrative DNA damage in animals and patients with inflammatory diseases in relation to inflammation-related carcinogenesis. Biol. Chem., 387, 365, 2006. 82. Ma, N. et al. Accumulation of 8-nitroguanine in human gastric epithelium induced by Helicobacter pylori infection. Biochem. Biophys. Res. Commun., 319, 506, 2004. 83. Pinlaor, S. et al. Nitrative and oxidative DNA damage in intrahepatic cholangiocarcinoma patients in relation to tumor invasion. World J. Gastroenterol., 11, 4644, 2005. 84. Srivatanakul, P. et al. Opisthorchis viverrini infestation and endogenous nitrosamines as risk factors for cholangiocarcinoma in Thailand. Int. J. Cancer, 48, 821, 1991. 85. Haswell-Elkins, M.R. et al. Liver fluke infection and cholangiocarcinoma: model of endogenous nitric oxide and extragastric nitrosation in human carcinogenesis. Mutat. Res., 305, 241, 1994. 86. Satarug, S. et al. Thiocyanate-independent nitrosation in humans with carcinogenic parasite infection. Carcinogenesis, 17, 1075, 1996. 87. Satarug, S. et al. Relationships between the synthesis of N-nitrosodimethylamine and immune responses to chronic infection with the carcinogenic parasite, Opisthorchis viverrini, in men. Carcinogenesis, 19, 485, 1998.

826 88. Satarug, S. et al. Induction of cytochrome P450 2A6 expression in humans by the carcinogenic parasite infection, Opisthorchiasis viverrini. Cancer Epidemiol. Biomarkers Prev., 5, 795, 1996. 89. Ohshima, H. et al. Increased nitrosamine and nitrate biosynthesis mediated by nitric oxide synthase induced in hamsters infected with liver fluke (Opisthorchis viverrini). Carcinogenesis, 15, 271, 1994. 90. Thanan, R. et al. Urinary 8-oxo-7,8-dihydro-2′-deoxyguanos­ ine in patients with parasite infection and effect of antiparasitic drug in relation to cholangiocarcinogenesis. Cancer Epidemiol. Biomarkers Prev., 17, 518, 2008. 91. Pinlaor, S. et al. Opisthorchis viverrini antigen induces the expression of Toll-like receptor 2 in macrophage RAW cell line. Int. J. Parasitol., 35, 591, 2005. 92. Pinlaor, S. et al. Repeated infection with Opisthorchis viverrini induces accumulation of 8-nitroguanine and 8-oxo-7,8dihydro-2′-deoxyguanine in the bile duct of hamsters via inducible nitric oxide synthase. Carcinogenesis, 25, 1535, 2004. 93. Thuwajit, C. et al. Increased cell proliferation of mouse fibroblast NIH-3T3 in vitro induced by excretory/secretory product(s) from Opisthorchis viverrini. Parasitology, 129, 455, 2004. 94. Pinlaor, S. et al. iNOS-dependent DNA damage via NF-kappaB expression in hamsters infected with Opisthorchis viverrini and its suppression by the antihelminthic drug praziquantel. Int. J. Cancer, 119, 1067, 2006. 95. Boonmars, T. et al. Apoptosis-related gene expression in hamster opisthorchiasis post praziquantel treatment. Parasitol. Res., 102, 447, 2008. 96. Maeda, H. Carcinogenesis via microbial infection. Gan To Kagaku Ryoho, 25, 1474, 1998. 97. Holzinger, F. et al. Mechanisms of biliary carcinogenesis: a pathogenetic multi-stage cascade towards cholangiocarcinoma. Ann. Oncol., 10 (Suppl. 4), 122, 1999. 98. Upatham, E.S. et al. Rate of re-infection by Opisthorchis viverrini in an endemic northeast Thai community after chemotherapy. Int. J. Parasitol., 18, 643, 1988. 99. Sornmani, S. et al. A pilot project for controlling O. viverrini infection in Nong Wai, Northeast Thailand, by applying praziquantel and other measures. Arzneimittelforschung, 34, 1231, 1984. 100. Saowakontha, S. et al. Field trials in the control of Opisthorchis viverrini with an integrated programme in endemic areas of northeast Thailand. Parasitology, 106, 283, 1993. 101. Hong, S. T. et al. The Kato-Katz method is reliable for diagnosis of Clonorchis sinensis infection. Diagn. Microbiol. Infect. Dis., 47, 345, 2003. 102. Viyanant, V. et al. A comparison of a modified quick-Kato technique and the Stoll dilution method for field examination for Opisthorchis viverrini eggs. J. Helminthol., 57, 191, 1983. 103. Radomyos, P. et al. Multi-infection with helminths in adults from northeast Thailand as determined by post-treatment fecal examination of adult worms. Trop. Med. Parasitol., 45, 133, 1994. 104. Radomyos, P., Bunnag, D. and Harinasuta, T. Haplorchis pumilio (Looss) infection in man in northeastern Thailand. Southeast Asian J. Trop. Med. Public Health, 14, 223, 1983. 105. Chai, J. Y. et al. High prevalence of liver and intestinal fluke infections among residents of Savannakhet Province in Laos. Korean J Parasitol, 45, 213, 2007.

Molecular Detection of Foodborne Pathogens 106. Poopyruchpong, N. et al. Diagnosis of opisthorchiasis by enzyme-linked immunosorbent assay using partially purified antigens. Asian Pac. J. Allergy Immunol., 8, 27, 1990. 107. Wongsaroj, T. et al. Affinity purified oval antigen for diagnosis of Opisthorchiasis viverrini. Asian Pac. J. Allergy Immunol., 19, 245, 2001. 108. Sirisinha, S. et al. Immunological analysis of Opisthorchis and Clonorchis antigens. J. Helminthol., 64, 133, 1990. 109. Sakolvaree, Y. et al. Parasites elicited cross-reacting antibodies to Opisthorchis viverrini. Asian Pac. J. Allergy Immunol., 15, 115, 1997. 110. Ruangsittichai, J. et al. Opisthorchis viverrini: identification of a glycine-tyrosine rich eggshell protein and its potential as a diagnostic tool for human opisthorchiasis. Int. J. Parasitol., 36, 1329, 2006. 111. Thammapalerd, N. et al. Detection of antibodies against Opisthorchis viverrini in patients before and after treatment with praziquantel. Southeast Asian J. Trop. Med. Public Health 19, 101, 1988. 112. Laha, T. et al. Gene discovery for the carcinogenic human liver fluke, Opisthorchis viverrini. BMC Genomics, 8, 189, 2007. 113. Sirisinha, S. et al. Evaluation of a monoclonal antibody-based enzyme linked immunosorbent assay for the diagnosis of Opisthorchis viverrini infection in an endemic area. Am. J. Trop. Med. Hyg., 52, 521, 1995. 114. Chaicumpa, W. et al. Specific monoclonal antibodies to Opisthorchis viverrini. Int. J. Parasitol., 21, 969, 1991. 115. Sirisinha, S. et al. Detection of Opisthorchis viverrini by monoclonal antibody-based ELISA and DNA hybridization. Am. J. Trop. Med. Hyg., 44, 140, 1991. 116. Wongratanacheewin, S. et al. Detection of Opisthorchis viverrini in human stool specimens by PCR. J. Clin. Microbiol., 40, 3879, 2002. 117. Wongratanacheewin, S. et al. Development of a PCR-based method for the detection of Opisthorchis viverrini in experimentally infected hamsters. Parasitology, 122, 175, 2001. 118. Stensvold, C.R. et al. Evaluation of PCR based coprodiagnosis of human opisthorchiasis. Acta Trop., 97, 26, 2006. 119. Duenngai, K. et al. Improvement of PCR for Detection of Opisthorchis viverrini DNA in Human Stool Samples. J. Clin. Microbiol, 46, 366, 2008. 120. Parvathi, A. et al. Development and evaluation of a polymerase chain reaction (PCR) assay for the detection of Opisthorchis viverrini in fish. Acta Trop., 107, 13, 2008. 121. Ando, K. et al. Nucleotide sequence of mitochondrial CO I and ribosomal ITS II genes of Opisthorchis viverrini in northeast Thailand. Southeast Asian J. Trop. Med. Public Health, 32 (Suppl. 2), 17, 2001. 122. Le, T.H. et al. Clonorchis sinensis and Opisthorchis viverrini: development of a mitochondrial-based multiplex PCR for their identification and discrimination. Exp. Parasitol., 112, 109, 2006. 123. Pauly, A. et al. Molecular characterization and differentiation of opisthorchiid trematodes of the species Opisthorchis felineus (Rivolta, 1884) and Metorchis bilis (Braun, 1790) using polymerase chain reaction. Parasitol. Res., 90, 409, 2003. 124. Ausubel, F.M., et al. Short Protocols in Molecular Biology, 3rd ed, John Wiley and Sons, 2. NewYork, 1995. 125. Sermswan, R. et al. Isolation and characterization of Opisthorchis viverrini specific DNA probe. Mol. Cell. Probes, 5, 399, 1991.

61 Paragonimus Kanwar Narain

Indian Council of Medical Research

Takeshi Agatsuma

Kochi Medical School

David Blair

James Cook University

Contents 61.1. Introduction..................................................................................................................................................................... 827 61.1.1. Classification and Morphology of Paragonimus Species................................................................................. 828 61.1.1.1. Paragonimus westermani (Kerbert, 1878) sensu lato...................................................................... 828 61.1.1.2. Paragonimus skrjabini (Chen, 1959) and P. miyazakii (Kamo et al., 1961)..................................... 829 61.1.1.3. Paragonimus heterotremus (Chen and Hsia, 1964).......................................................................... 829 61.1.1.4. Paragonimus africanus (Voelker and Vogel, 1965)......................................................................... 829 61.1.1.5. Paragonimus uterobilateralis (Voelker and Vogel, 1965)................................................................ 829 61.1.1.6. Paragonimus mexicanus (Miyazaki and Ishii, 1968)....................................................................... 829 61.1.1.7. Paragonimus kellicotti (Ward, 1908)............................................................................................... 829 61.1.2. Biology and Pathogenesis.................................................................................................................................. 829 61.1.3. Methods for Diagnosing Infection due to Paragonimus Species...................................................................... 830 61.1.3.1. Immunological Techniques............................................................................................................... 830 61.1.3.2. Molecular Techniques....................................................................................................................... 832 61.2. Methods........................................................................................................................................................................... 832 61.2.1. Sample Collection and Preparation................................................................................................................... 832 61.2.1.1. Sample Collection............................................................................................................................. 832 61.2.1.2. DNA Extraction from Eggs, Metacercariae, and Adult Worms....................................................... 833 61.2.2. Detection Procedures........................................................................................................................................ 833 61.3. Conclusions and Future Perspectives.............................................................................................................................. 835 References.................................................................................................................................................................................. 835

61.1  Introduction Paragonimiasis or endemic hemoptysis is an important disease of man and other mammals caused by several trematode species belonging to the genus Paragonimus.1,2 Typically, paragonimiasis is a disease of the lungs and pleural cavity but extra-pulmonary paragonimiasis is also an important clinical manifestation. Lung-flukes utilize a mollusk as a first intermediate host and a freshwater crab or crayfish as a second intermediate host.3 Consumption of raw, undercooked or pickled freshwater crab or crayfish introduces lung-flukes into the human body.2 Similarly, raw or undercooked meat of paratenic hosts such as boar, bear, wild pig or rat, in the muscles of which juvenile worms can survive for years, is also an important source of human infection.4 Paragonimiasis is a neglected disease that has received relatively little attention from public health authorities. It has

been estimated that around 293 million people are at risk and several million are infected worldwide.5 However, this may be an underestimate as there are still many places where the disease burden has yet to be assessed. In recent times there has been increased recognition of the public health importance of paragonimiasis and other foodborne trematodiases.6 In the case of paragonimiasis, this resurgence of interest is partly due to the common diagnostic confusion of paragonimiasis with tuberculosis. Symptoms of the former closely mimic those of tuberculosis, frequently leading to inappropriate treatment being administered especially in areas where both tuberculosis and paragonimiasis co-occur and are overlapping public health issues.7 Failure of patients to respond to treatment for tuberculosis may lead to inflated estimates of the prevalence of multi-drug resistant tuberculosis and have other far-reaching health implications.8–12 Interest in 827

828

Paragonimus species outside endemic areas is also increasing because of the risk of infection through consumption of crustaceans traded far from their point of origin in today’s globalized food supply. Traditionally, morphological criteria were used for identification and taxonomy of lung-flukes. However, the explosion of molecular techniques in recent years has led to the development of better diagnostic tools and a better understanding of lung-fluke taxonomy. Nevertheless, there is still confusion regarding the taxonomic status of a number of species. Molecular characterization of lung-flukes will improve our understanding of evolution and epidemiology of paragonimiasis and our capacity to control this foodborne disease.

61.1.1 Classification and Morphology of Paragonimus Species Taxonomic studies of lung-flukes (Order Plagiorchiida, Suborder Xiphidiata, Superfamily Gorgoderoidea, Family Paragonimidae13 can be broadly divided into two phases. In the first phase (up to the late 1990s), much emphasis was laid on the gross morphological features (Adult worms: body shape and size, relative sizes of suckers, patterns of lobation of the ovary and testes, arrangement of cuticular spines. Metacercariae: presence or absence, number, and thickness of cyst walls, relative diameter of suckers, the anterior extent of the excretory bladder, presence, and length of a stylet on the oral sucker, presence of colored granules in the body, number of flame cells, body spination and arrangement of papillae around suckers. Eggs: size, egg shell sculpturing). During the second phase (from the early 1990s), DNA sequence data from nuclear and mitochondrial genes were increasingly used to aid taxonomy and to infer phylogenetic relationships of lung-flukes. The genus Paragonimus (Braun, 1899) was erected to include the type species P. westermani (Kerbert, 1878). Subsequently, two additional genera of lung-flukes were created: Pagumogonimus (Chen, 1963) and Euparagonimus (Chen, 1963), based on morphological characteristics of adult flukes and metacercariae.1 However, there is considerable plasticity of morphological characters, together with apparent homoplasy of most of these characters, making for taxonomic confusion. DNA sequence data and phylogenetic analysis showed that the two genera Pagumogonimus and Euparagonimus are artificial and thus untenable.14 More than 50 nominal species of the genus Paragonimus have been reported from Asia, Africa, and the Americas,1 a number difficult to justify based on the rather few morphological characters available. DNA sequence data, both mitochondrial DNA and nuclear ribosomal DNA, are now being used extensively to characterize lung-flukes and to resolve some questions of their taxonomy and evolution.2,14–25 Such molecular studies have demonstrated that some nominal species of lung-flukes are best regarded as species complexes, a topic discussed further below. Even though controversies remain regarding the number of valid species in this genus Paragonimus it is clear that at

Molecular Detection of Foodborne Pathogens

least seven species (or species-complexes) can cause human infection.2 The most important of these are listed below. 61.1.1.1 Paragonimus westermani (Kerbert, 1878) sensu lato Probable synonyms:1 P. ringeri (Cobbold, 1880) (Ward and Hirsch, 1915); P. pulmonalis (Baelz, 1880) (Miyazaki, 1978); P. pulmonis (Suga, 1883); P. macacae (Sandosham, 1953); P. edwardsi (Gulati, 1926); P. asymmetricus (Chen, 1977); P. filipinus (Miyazaki, 1978); P. philippinensis (Ito, Yokogawa, Araki, and Kobayashi, 1978). P. westermani was discovered by Kerbert (1878) in the lungs of a Bengal tiger that had died in the Amsterdam Zoological Gardens. The tiger is presumed to have come from India, but its origins cannot be confirmed.1 Subsequently, the species has been reported from many taxa of mammals.1 What is generally called P. westermani has a wide geographical distribution, extending from eastern Asia (China, Taiwan, Korea, Japan, Southeast Siberia); South East Asia (Philippines, Malaysia, Thailand, Cambodia, Laos, Vietnam), South Asia (Sri Lanka, India, Nepal, Pakistan). Endemic human paragonimiasis due to P. westermani is recorded only from East Asia (Japan, Korea, China, Taiwan), and the Philippines. A single apparently autochthonous human case from Papua New Guinea was possibly due to this species.26 Populations of the parasite from elsewhere in its broad range are not known to infect humans. The body of adult P. westermani is discoidal or slightly elongated when compressed, the ovary is distinctly six-lobed, and the testes also subdivided into five or six lobes and similar in size to the ovary. The oral sucker is slightly larger than the ventral. Metacercariae are typically spherical in shape and possess two cyst walls. The inner wall is thick, although some populations have a thin wall. Metacercariae of P. westermani from India and Sri Lanka are oval in shape and do not possess thick inner cyst walls.21–23 As might be suspected from the above, P. westermani is probably a species-complex rather than a single species. This is also evident from geographical variation in biological features such as host-specificity and suggested by molecular phylogenies. Genetic studies commencing about 20 years ago have demonstrated considerable geographical variation among P. westermani-like lung-flukes. An early populationgenetic analysis using eight enzyme loci demonstrated that a Philippine population of P. westermani was genetically quite different from populations from Japan and Taiwan.27 Similarly, worms from a Malaysian population were found to be distinct from those from Japan and the Philippines in a study utilizing 20 enzyme loci.28 Based on this and later molecular sequence data, it is clear that nomenclatural difficulties remain with P. westermani.21 Further complicating understanding of the P. westermani group is the fact that polyploid forms, usually triploids, can occur, often sympatric with diploids, in Japan, Korea, China, and Taiwan. Whereas diploid individuals require a mate for exchange of sperm before they can produce viable eggs,

829

Paragonimus

triploid worms are parthenogenetic, larger than diploids and able to produce viable eggs without a mate.1 61.1.1.2 Paragonimus skrjabini (Chen, 1959) and P. miyazakii (Kamo et al., 1961) Probable synonyms:1,20 P. miyazakii (Kamo, Nishida, Hatsushika, and Tomimura, 1961) (reduced to subspecific status); P. szechuanensis (Chung and Tsao, 1962); P. hueitungensis (Chung, Hsu, Ho, Kao, Shao, Chiu, Pi, Liu, Ouyang, Shen, Yi, and Yao, 1975); P. veocularis (Chen and Li, 1979). P. skrjabini represents another species-complex that has recently been subject to comprehensive morphological and molecular analysis. Nominal P. skrjabini from Fujian, China is phylogenetically very close to P. miyazakii from Japan and consequently this population was placed with Japanese populations as P. skrjabini miyazakii.20 It was also proposed that populations from other parts of China (Guangdong, Sichuan, and Hubei Provinces and Guangxi Zhuang Autonomous Region) should be named P. skrjabini skrjabini. The known geographical distribution of the P. skrjabini species complex is restricted to China, Japan, Thailand, Vietnam, and northeastern India.20,29,30 The natural definitive hosts of members of this species-complex include many small carnivorous or omnivorous mammals.1 Members of the species-complex have distinctly elongated bodies (length:width ratio between 1.8 and 2.9). The ventral sucker is very slightly larger than the oral sucker and is about one-third from anterior end of body. Testes are typically larger and show a much greater degree of variation in size and lobation than is seen in members of the P. westermani complex. The ovary is profusely branched. The inner cyst of the metacercaria is thick and the juvenile worm occupies the entire internal cyst space, but geographical variation occurs within the complex.20 61.1.1.3  P  aragonimus heterotremus (Chen and Hsia, 1964) Probable synonym:1 Paragonimus tuanshanensis (Chung, Ho, Cheng, and Tsao, 1964). P. heterotremus occurs in eastern Asia, from NE India to southern China, as well as Vietnam, Thailand, and Laos.31–34 Adults have been reported from naturally infected dogs, cats, and rodents.1 The species is morphologically distinctive, both as an adult and a metacercaria. An unusual feature of the adult fluke is the size difference between the suckers, a feature it shares with P. africanus. In most other Paragonimus species, the two suckers are approximately equal in size: in P. heterotremus the ventral sucker is about half the diameter of the oral. The metacercaria of P. heterotremus is easily identifiable because of its small size and the thickening of the inner cyst wall at both poles.3 61.1.1.4 Paragonimus africanus (Voelker and Vogel, 1965) P. africanus was first found in mongooses from Lower Bakossi, West Cameroon and is now known from Equatorial Guinea, Cameroon, Nigeria, and Ivory Coast.35–39 Natural

definitive hosts include lorids, cercopithecids, herpestids, and viverrids.1 In the adult flukes, the oral sucker is distinctly larger than the ventral sucker. The ovary is delicately branched whereas the testes are highly branched and significantly larger than the ovary. Metacercariae are spherical in shape with two cyst walls. 61.1.1.5 Paragonimus uterobilateralis (Voelker and Vogel, 1965) P. uterobilateralis was first found in mongooses from Cameroon and is now known from Gabon, Cameroon, Nigeria, Liberia.40–45 Natural definitive hosts include canids, herpestids, viverrids, and mustelids.1 In the adult flukes, oral, and ventral suckers are similar in size. Cuticular spines are arranged singly. The ovary is delicately branched whereas the testes are moderately branched and larger than the ovary. Metacercariae are rounded with a thin single cyst wall. 61.1.1.6 Paragonimus mexicanus (Miyazaki and Ishii, 1968) Probable synonyms:1 P. peruvianus (Miyazaki, Ibañéz, and Miranda, 1969); P. ecuadoriensis (Voelker and Arzube, 1979). This species, widespread between Mexico and Peru, is the only species known to cause paragonimiasis in Latin America.3 Natural mammalian definitive hosts include didelphids, cebids, canids, felids, mustelids, procyonids, and suids.1 Adult worms are somewhat elongated, possess strongly lobed ovary and testes, and the oral sucker is somewhat larger than the ventral. Unlike any of the other species listed here, the egg-shell is pitted in a distinctive manner. Metacercariae are found free in the tissues of crab intermediate hosts. 61.1.1.7 Paragonimus kellicotti (Ward, 1908) This species is found primarily in the central and eastern states of the United States and adjacent parts of southern Canada. Adult worms have been reported from a number of mammal species including domestic pigs, dogs, and cats. Natural hosts include small carnivorous or omnivorous mammals.1 This species was originally confused with P. westermani, but adults of P. kellicotti have gonads that are more branched than in P. westermani. Metacercariae are spherical and possess two cyst walls. They occur in freshwater crayfish.

61.1.2  Biology and Pathogenesis Lung-flukes have a three-host life cycle. Typically, maturing worms meet potential mates in the thoracic cavity of their natural mammalian host. Pairs of worms then move into the lung parenchyma where a fibrous cyst develops around them. After exchange of sperm, each of the hermaphroditic worms produces huge numbers of eggs that escape from the lung cyst into small bronchioles and eventually to the external environment via sputum or feces. A ciliated miracidium escapes from the egg after hatching in freshwater and must penetrate a suitable mollusk, a first intermediate host, within hours.1 Cycles of asexual reproduction within the mollusk eventually yield large numbers of cercariae that escape from the snail

830

Molecular Detection of Foodborne Pathogens

and encyst within a freshwater crab or crayfish as metacercariae.1 These, emerging as juvenile worms from their cysts in the gut of mammals eating raw crab or crayfish, must migrate through the tissues of the mammal into the pleural spaces to continue the life cycle. If the crab or crayfish host is eaten by a mammal that is not suitable for maturation of the worms, juvenile lung-flukes will remain alive within the muscles, potentially for long periods of time, and can mature if their paratenic host is finally eaten by a suitable mammal species. A range of symptoms and signs have been reported in human paragonimiasis.46 Pulmonary paragonimiasis is the best known clinical manifestation. As in their natural mammal hosts, pairs of adult worms come to reside in a fibrous cyst within the lungs of humans and produce eggs. Worm pairs can survive for some years. Symptoms of established infection include cough and the production of sputum containing blood and brown parasite eggs. Symptoms can be severe in heavy infections and seem to have been particularly pronounced in the few known human cases of infection with P. kellicotti in North America.47,48 Pulmonary forms of the disease are typical in people infected with P. westermani, P. heterotremus, P. africanus, P. uterobilateralis, P. mexicanus and P. kellicotti. Pleural lesions, such as pleural effusion, pneumothorax, and pleural thickening, can be associated with pulmonary paragonimiasis. P. skrjabini miyazakii, the representative of the P. skrjabini group endemic to Japan typically produces pleural and ectopic lesions, but can also cause pulmonary disease.2 Adult and juvenile lung-flukes may wander far from the lungs and thoracic region. Particularly unpleasant symptoms can result from the presence of worms in the brain. Other sites from which worms have been recorded include the

skin, liver, abdominal organs, gonads, and genitalia, eye, and spinal cord.2,46

61.1.3 Methods for Diagnosing Infection due to Paragonimus Species Definitive diagnosis of paragonimiasis depends upon finding the characteristic operculated eggs in the sputum of patients. However, eggs may not be detected in the sputum of infected persons because of light infections (low egg output) or intermittent egg production. In such situations, up to seven repeated sputum examinations have been recommended to maximize the chance of correct diagnosis in suspected patients.7 Examination can be accomplished by simple microscopy of a wet sputum smear. Concentration methods have also been used to achieve a higher sensitivity for detection of pulmonary paragonimiasis. Sputum is liquefied by adding double the volume of 4% caustic soda followed by mixing and incubation at 37°C for 20–30 min. Finally, the sample is centrifuged and the sediment examined under the microscope. In case of children below 10 years it has been recommended to examine stool samples for Paragonimus eggs as children often swallow the sputum and consequently eggs are present in the feces.7 In some situations more invasive laboratory procedures such as bronchoscopic washings, percutaneous lung aspiration, and fine needle biopsy may be required for detection of lung-fluke eggs to establish the correct diagnosis (Table 61.1).49 61.1.3.1  Immunological Techniques Although the detection of eggs is confirmatory of paragonimiasis, nevertheless eggs are difficult to detect in extrapulmonary paragonimiasis. Indeed, the worms in such cases may not even reach maturity and hence may not produce

Table 61.1 Summary of Diagnostic Methods for Paragonimiasis Diagnostic Methods

Paragonimus ova can be found in sputum or feces or occasionally in bronchoscopic washings, biopsy specimens, fine needle aspirate, or pleural effusion of the definitive host by microscopy.74–77

Remarks Parasitological Diagnosis • Identification of eggs of different species of Paragonimus by microscopy is often difficult and problematic. In stool and other clinical samples, diagnostic confusion can arise because of the presence of eggs of other species of trematodes (e.g., Achillurbainia congolensis78) or operculate cestode eggs (e.g., Diphyllobothrium latum). Note: Acid-fast staining of a sputum smear for detection of Mycobacterium tuberculosis often destroys Paragonimus eggs. So separate slides should be prepared for examination of sputum samples in such cases.

PCR amplification and gene sequencing for identification of lung fluke species causing infection in man or other animals. PCR-RFLP for molecular discrimination of Paragonimus causing infection in man or animals.

Molecular Diagnosis • For molecular identification, internal transcribed spacer 2 (ITS2) of the nuclear ribosomal RNA repeat unit, and mitochondrial cytochrome c oxidase subunit 1 (cox1) can be sequenced for species identification (e.g., detection of P. westermani eggs from human sputum samples.31,68 • For discrimination between P. westermani and P. miyazakii restriction digestion of ITS2 PCR product.69 • For discrimination of P. westermani and P. heterotremus.70 (Continued)

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Table 61.1  (Continued) Diagnostic methods Species-specific PCR.

Multiplex PCR in combination with RFLP.

Species-specific DNA probe.

Monoclonal antibody-based antigen detection assay.

Locally developed ELISAs using crude worm extracts or excretory/secretary antigens have been developed. Recombinant yolk ferritin-based ELISA.

Western blotting for diagnosis of paragonimiasis.

Remarks • Identification of metacercariae of P. westermani and P. heterotremus.70 For P. westermani, a specific forward primer (PwTF1-5′-GTT CAT GTT GCG CGT GGT CTG CGT TC-3′) and a “universal” reverse primer A28 (5′-GGG ATC CTG GTT AGT TTC TTT TCC TCC GC-3′) are used and for P. heterotremus, a specific forward primer (PhTF1-5′-TTC CCC AAC GTG GCC TTG TGT-3′) and reverse primer A28 are used. Although the above method was developed for metacercarial identification these methods can also be used to differentiate other life cycle stages of the flukes. • For detection of P. heterotremus eggs in experimentally infected cats.79 Species-specific primers: forward primer (5′-GGG TAG CAC CAT CTC GT-3′) and reverse primer (5′-GAT CAC GAG GAC CCC AT-3′). • For discrimination of individual metacercariae of five different lung fluke species (P. westermani, P. heterotremus, P. siamensis, P. bankokensis and P. harinasutai) occurring in Thailand.80 The above method was developed for Thai strains of lung flukes only and therefore needs to be validated for other strains of lung flukes which originate from different geographical areas. • For detection of P. heterotremus eggs in stool samples.67 A species-specific DNA probe 1473 bp long (GenBank accession no. AZ254640). This probe has high specificity for P. heterotremus and does not show cross-reactions with genomes of related parasites, but its use is limited because of low sensitivity. Antigen Detection • The performance of Dot-ELISA for diagnosis of active paragonimiasis was evaluated to detect species- and stage-specific antigens of Paragonimus westermani. The monoclonal antibody MAbA3D12D3 reacted with a polypeptide determinant of adult worm antigen and the monoclonal antibody MAb 2G2B3 reacts with the carbohydrate epitope of a metacercarial antigen of P. westermani.81 Also demonstrated was the sensitivity of the antigen detection assay to monitor the efficacy of praziquantel for treatment of Paragonimus westermani infections.82 Antibody Detection The utility of somatic and excretory/secretory antigens for detection of antibodies against human paragonimiasis was demonstrated.9,56 Compared with somatic antigens the excretory/ secretory antigens were shown to be highly specific for paragonimiasis. • A sero-diagnostic test based on detection of antibodies against recombinant P. westermani yolk ferritin is useful in early stages of paragonimiasis.83 • A 322 amino acid based recombinant protein, mostly distributed in eggs and uterus of P. westermani, was evaluated for serodiagnosis of paragonimiasis.84 The sensitivity of ELISA based using this recombinant protein was 90.2% with a specificity of 100%. • In an immunoblot assay using Paragonimus westermani Chaffee antigen, antibody reactivity to an 8-kDa band was shown to be diagnostic with a sensitivity of 96% and specificity of 99%.62 • A 31.5-kDa band was reactive in an immunoblot assay based on P. heterotremus excretory-secretory products. The sensitivity and specificity for diagnosis of human P. heterotremus infection was 97% and 100%, respectively.85 • An EITB assay based on antigens from P. heterotremus adult worms revealed bands (35, 33 and 32.5 kDa) diagnostic of P. heterotremus infection and bands (34.5, 34, 33, and 32.5 kDa) diagnostic of P. westermani nfection.66 • An immunoblot assay using a recombinant 28-kDa Cruzipain-like cysteine protease was useful for sero-diagnosis of active paragonimiasis with a sensitivity of 86.2% and specificity of 98%.86

eggs at all. Therefore, immunological tests are valuable. Immunodiagnostic tests have also been successfully used for epidemiological surveys and studies involving post treatment evaluation (Table 61.1). Intradermal test: The intradermal test is an old diagnostic test based on cellular responses to injected antigens as an

indicator of infection. An extract from lung-flukes is injected into the skin on the volar aspect of the forearm and the size of the initial wheal noted.50–52 The test is considered positive if the diameter of the wheal has increased by more than 5 mm after some 15 min.52 Many different antigen extracts have been used.51,52 Side effects are relatively rare. Disadvantages of the

832

intradermal test include the high frequency of false positives, but false negatives are rare. Another problem is that the test can remain positive for long periods after the death of all worms in a patient. ELISA: Enzyme linked immunosorbent assay (ELISA) is a reliable immunological test widely used for detection of antibodies in sera, pleural effusions and cerebrospinal fluid. Conventionally, the ELISA test uses a multi-well plate to provide a solid surface support on which an antigen or an antibody can be bound for subsequent reactions.1,2 Researchers from Thailand and China were among the first to use ELISA for diagnosis of paragonimiasis.53,54 Crude somatic antigen can be a reliable reagent for sero-diagnosis of paragonimiasis in ELISA, regardless of the causative species.55 Crude antigen from P. westermani, P. miyazakii, P. heterotremus, and P. siamensis could all be used for the detection of P. westermani infection.55 Despite such results using somatic antigens, it has been shown that excretory and secretory (ES) antigens of adult lung-flukes are highly specific as compared to somatic antigens.9,56 A number of studies have focused on methods for purifying and preparing various types of antigen to improve specificity in ELISA. These methods include: preparative isoelectric focusing for partial purification of adult somatic antigens of P. heterotremus,57 use of affinity-purified ES antigen,58 development of a recombinant P. westermani yolk ferritin antigen found to be highly reliable for serodiagnosis of P. westermani in early and chronic stages.59 Anti-Paragonimus antibodies (IgG) persist for 4–18 months after praziquental treatment, suggesting that an IgG ELISA test should be conducted 18 months post treatment to confirm cure.60 For diagnosis of active P. westermani infection, an antigen detection assay has been developed using monoclonal antibodies for capture of circulatory antigens from sera of infected patient.61 Immunoblotting: Immunoblot assays can be sensitive and specific for diagnosis of paragonimiasis.62 In one study, P. westermani Chaffee antigen for SDS PAGE and western blot analysis were used. Out of 45 serum samples from confirmed paragonimiasis cases, 43 reacted with an 8-kDa band.62 The sensitivity and specificity of this immunoblot test was 96% and 99%, respectively.62 Other workers have identified different antigenic proteins (27 kDa;63 27 kDa and 33–37 kDa;64,65 23, 30, and 46 kDa60,65) from P. westermani, which react with sera of infected patients. These differences may be due to variation in antigen preparation, SDS-PAGE, etc.60 Differential diagnosis between P. heterotremus and P. westermani was possible based on ether-extracted antigens from adult worms of P. heterotremus. Reactive bands of 35, 33, and 32.5 kDa in the immunoblot reaction were diagnostic of P. heterotremus infections whereas the 35 kDa band was not reactive in P. westermani infection.66 61.1.3.2  Molecular Techniques DNA probes: A highly sensitive and specific DNA probe, 1500 bp long, was characterized for P. heterotremus.67

Molecular Detection of Foodborne Pathogens

The labeled probe could detect DNA from as few as five eggs with 100% specificity but low sensitivity.67 PCR: Parasitological identification of Paragonimus species is mostly based on microscopic observations of eggs in sputum samples. Unfortunately, eggs from different species show very subtle differences in their morphological characteristics including shape and size. However, it has proved possible to amplify and sequence DNA from a small number of eggs and thus confirm the identity of the lung-fluke.31,68 In one case, the results were unexpected: DNA sequences from eggs demonstrated that human infection in Arunachal Pradesh, India is due to P. heterotremus, and not P. westermani as had been assumed.31 Several varieties and combinations of PCR-RFLP (restriction fragment length polymorphism) and species-specific PCR amplification have been used to distinguish between species. Amplification of ITS2 using conserved primers and subsequent digestion with the restriction endonuclease SnaB1 yielded two bands in the case of P. westermani, but a single (undigested) band in the case of P. miyazakii.69 Conversely, there is no cut site for BssSI in the ITS2 of P. westermani but a single cut site exists in P. miyazakii. This can provide an easy way to distinguish between metacercariae of these two medically important species that can co-occur within the same crab in Japan. Similar approaches permit discrimination between metacercariae of P. heterotremus and P. westermani in Thailand (although it should be noted that these are very easy to discriminate on morphology alone).70 It is also feasible to design primer pairs that will amplify a target region (typically ITS2) from one lung-fluke species, but not from others (Table 61.1).70

61.2  Methods Various diagnostic strategies and current molecular approaches to the identification of lung-flukes are discussed. Table 61.1 summarizes the current techniques available for diagnosis of paragonimiasis

61.2.1 Sample Collection and Preparation 61.2.1.1 Sample Collection The traditional method of detection and collection of lungfluke metacercariae is by microscopical examination of the crab and crayfishes. Gills, heart, hepatopancreas, thoracic muscles and muscles of the appendages should be examined. Compression of these parts of the crab between two glass slides facilitates the detection of metacercariae. Some workers use acid-pepsin solution (artificial digestive juice) for digestion of the host tissue for recovery of metacercariae.71 In brief, the crab tissues are incubated in a shaking water bath for 1½–2 h at 37°C and subsequently, the digested material is passed through two layers of wet gauze. The digested material is allowed to sediment in a tall glass cylinder containing physiological saline and the sediment is washed a number of times using physiological saline. The sediment

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Paragonimus

is finally examined under a stereomicroscope for detection of lung-fluke metacercariae. Identification to species can be done using morphological features72 or molecular data. Adult worms can be raised in experimental animals for morphological identification of species but this is time consuming, expensive, and ethically questionable. Furthermore, morphological features are of little help in resolving some closelyrelated species.20 61.2.1.2 DNA Extraction from Eggs, Metacercariae, and Adult Worms We have found E.Z.N.A. Mollusk DNA Isolation Kit (Omega Biotek) efficient for extraction of genomic DNA from different stages of lung-flukes. This kit is also efficient for recovery of genomic DNA from other flatworms and other invertebrate tissue rich in mucopolysaccaride and is suitable for frozen tissues or samples preserved in alcohol. Reagents: (i) E.Z.N.A. Mollusk DNA Isolation Kit and the reagents include with the kit; (ii) Proteinase K (stock, 20 mg/ml); (iii) chloroform: isoamyl alcohol (24:1) mixture; (iv) RNase A (20 µg/ml); (v) absolute ethanol. Equipment: (i) sterile Eppendorf centrifuge tubes (1.5 ml or 2 ml) and matching sterile polypropylene pestles for grinding; (ii) SpeedVac vacuum concentrator (Savant) or similar; (iii) vortexer; (iv) incubator or water bath capable of reaching at least 70°C; (v) microfuge capable of spinning Eppendorf tubes at 15,000 × g. Procedure: (1) Remove alcohol from samples by drying in SpeedVac for 10 min. (2) Grind the dried material in a sterile Eppendorf centrifuge tube with the help of a sterile polypropylene pestle. (3) Add 350–550 µl buffer (ML1) followed by 25 µl proteinase K. Mix in vortexer and incubate at 60°C for minimum 1–5 h (or until the entire sample become solubilized). (4) To the lysate add 350 µl chloroform: isoamyl alcohol (24:1) and mix in vortexer. Centrifuge at 10,000 × g for 2 min at room temperature. Transfer the upper aqueous phase into a clean 1.5 ml microfuge tube. (5) Add one volume of buffer MBL followed by 5 µl of RNase A, vortex for 15 sec. Incubate at 70°C for 10 min. (6) Add one volume of absolute ethanol (room temperature) and vortex briefly. (7) Take 750 µl of the mixture, including any precipitate, to a spin column assembled in a 2-ml ­collecting tube (supplied with the DNA isolation kit). Centrifuge at 10,000 × g for 1 min at room temperature and ­discard flow-through liquid and reuse collection tube. (8) Repeat step 7 for remaining mixture from step 6. (9) Place the column into a new 2-ml collection tube (supplied with the DNA isolation kit) and wash by









adding 500 µl HB buffer. Centrifuge at 10,000 × g for 30 sec. Discard collection tube along with the flow-through. (10) Place column into a new 2 ml collecting tube (supplied) and wash with the help of 700 µl DNA buffer diluted with absolute ethanol. Centrifuge at 10,000 × g for 1 min. Discard flow-through but reuse collection tube for next step. (11) Repeat step 10 with a second 700 µl DNA buffer diluted with ethanol. Discard liquid and re-insert the column to the empty collecting tube, centrifuge the column at 15,000 × g for 2 min at room temperature to remove traces of ethanol. (12) Place column into a clean 1.5-ml microfuge tube. Add 50–100 µl of elution buffer (or 10 mM Tris buffer, pH 9.0) preheated to 60–70°C directly onto the supplied HiBind spin column matrix. Then allow to soak for 2 min at room temperature. Centrifuge at 10,000 × g for 1 min to elute DNA. (13) Repeat elution step with a second 50–100 µl of elution buffer.

Note: (i) For sample collection and preservation it is best to store the sample at –80°C. Ethanol is also good for sample preservation but DNA in ethanol-preserved samples may degrade due to acid formation. (ii) There are other alternative methods for genomic DNA isolation from platyhelminth samples. The cheapest and simple method is Chelex extraction combined with proteinase K. This procedure requires incubation of the samples in the chelex/proteinase K mixture for 12–24 h at 55°C and requires the additional steps of heating the samples at 95°C for 20 min to inactivate proteinase K, followed by centrifugation. 1–2 µl of supernatant can be used for PCR.

61.2.2  Detection Procedures Principle: Use of specific primer pairs facilitates amplification of targeted regions of the genome of organisms of interest. If the PCR is successful, amplification products can be visualized on an agarose gel and their length checked against that expected. Subsequently, PCR products can be sequenced for confirmation of identity by comparison with available data. Use of both a nuclear region (e.g., ITS2) and a mitochondrial gene (e.g., cox1) is common when identification to species/subspecies/geographic strain is required. For amplification of the ITS2 region of Paragonimus species, the following pair of primers can be used: forward primer (3S) (5′-GGT ACC GGT GGA TCA CTC GGC TCG TG-3′) and reverse primer (A28) (5′-GGG ATC CTG GTT AGT TTC TTT TCC TCC GC-3′).18 For amplification of a portion of the cox1 gene, the following pair of primers can be used: forward primer (JB3) (5′-TTT TTT GGG CAT CCT GAG GTT TTA-3′) and reverse primer (JB4.5) (5′-TAA AGA AAG AAC ATA ATG AAA ATG-3′).73

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Molecular Detection of Foodborne Pathogens

Reagents for PCR: (i) primers as above; (ii) PCR mixture according to the following table: Reagents for Preparation of PCR Mixture

Volume Required for One 50 µl Reaction

1. 10 × Buffer (Mg +  + free) 2. dNTPs (stock: 10 mM each) 5. Forward primer (10 pmol/µl) 6. Primer 28A (reverse, 10 pmol/µl) 5. Taq DNA polymerase (5 units/µl) 6. BSA solution (10 mg/ml) 7. MgCl2 (25 mM) 8. Template DNA 9. Milli-Q water

  5.0 µl   1.0 µl   3.0 µl   3.0 µl 0.5–1 µl   0.5 µl   4.0 µl   2.0 µl To total volume of 50 µl

Other reagents: (i) TBE buffer (10 × stock): 1 M Tris, 1 M Boric acid, 50 mM EDTA (pH 8.0–8.3); (ii) agarose (such as NuSieve or MetaPhor); (iii) ethidium bromide solution (stock: 10 mg/ml in H2O: wrap container in aluminum foil and store in dark at 4°C); (iv) loading dye (2 ×); (v) Milli-Q water or autoclaved double distilled H2O. Equipment: (i) Thermocycler with heated lid; (ii) sterile powder-free gloves; (iii) sterile 1.5 ml Eppendorf tubes; (iv) 0.2 ml thin-walled PCR tubes; (v) micropipettes capable of handling appropriate volumes; (vi) sterile tips for micropipettes; (vii) sterile powder-free gloves; (viii) micropipettes capable of handling appropriate volumes; (ix) sterile tips for micropipettes; (x) gel tank, gel tray, combs, and power supply for agarose electrophoresis; (xi) UV transilluminator and camera or other gel-documentation system; (xii) scalpel or similar for band excision. (i) PCR amplification: (1) Remove the reagents required for PCR from the freezer and allow to thaw. Mix well and spin down. Prepare PCR mixture (50 μl) as outlined above. (2a) (ITS2 cycling conditions). Perform PCR amplification in a thermocycler for one cycle of 94°C for 3 min; 31 cycles of 94°C for 30 sec, 50°C for 30 sec, and 72°C for 30 sec; and one cycle of 72°C for 5 min. (2b) (cox1 cycling conditions). Perform PCR amplification in a thermocycler for one cycle of 94°C for 3 min; 35 cycles of 94°C for 30 sec, 48°C for 30 sec, and 72°C for 30 sec; and one cycle of 72°C for 5 min. (3) After completion of the PCR, mix 5 μl of PCR mixture with 2 μl of loading buffer and separate on a 2% agarose gel in a TBE buffer containing ethidium bromide (0.5 μg/ml). Run the gel for 30–45 min at 100 V. Examine the agarose gel under UV light to detect the presence of amplified PCR product of desired size. (ii) Purification of PCR fragment from gel: (1) Electrophorese the PCR product in a 1.2% agarose gel (containing 0.5 µg/ml ethidium bromide) in 1 × modified TAE buffer (prepared from 50 × stock).

(2) Locate the required DNA band in the gel using a long-wavelength UV lamp or a transilluminator. Cut and excise the gel slice containing the DNA (gel slice must be under 100 µl in volume or 100 mg in mass). (3) Place the gel slice in the Gel Nebulizer and seal the device with the cap attached to the vial (Geneclean kit, Q-BIO Gene, Carlsbad, CA). (4) Spin for 10 min at 5,000 × g. The agarose is captured in the sample filter cup. The DNA passes (in TAE buffer) through the sample filter cup and is collected in the filtrate vial. (iii) Cycle sequencing: (1) Thaw BigDye sequencing reagents on ice. Mix well and spin down. (2) Prepare the reaction mixture for cycle sequencing according to the recipe given below:

Reagents (1) Ready reaction mix (2) BigDye sequence buffer (v3.1) (3) *Primer (0.8 pmol/µl) (4) Template (Gene clean product) (5) Milli-Q water

Volume (For 20 µl of Reaction Volume) 4.0 µl 2.0 µl 4.0 µl (3.2 pmol) 1–3 µl To total volume of 20 µl

Note: Only one primer is included in any given reaction mix in cycle sequencing.

(3) For cycle sequencing, the conditions are one cycle of 96°C for 1 min; 25 cycles of 96°C for 30 sec, 50°C for 30 sec and 60°C for 4 min. The reaction products can be stored at -40°C until ready to purify. (iv) Ethanol precipitation of cycle sequencing product: (1) If held at -40°C, take out and thaw out, mix well and spin down. (2) Add 2.5 µl 125 mM EDTA. (3) Add 2.5 µl 3 M Na-acetate. (4) Add 50 µl of ice-cold ethanol (100%). (5) Mix by tapping. (6) Incubate at room temperature for 15 min. (7) Centrifuge at 14,000 × g for 20 min (4°C). (8) Discard the supernatant by inverting PCR tube into 1.5-ml Eppendorf tube and very briefly spinning at 150 × g. [Be very careful not to dislodge the pellet of extension products]. (9) Add 70% ice-cold ethanol (150 µl) and spin at 14,000 × g for 20 min. (10) Discard supernatant as in step 8 using fresh Eppendorf tubes. (11) Repeat steps 9 and 10. (12) Vacuum dry for 10 min (in DNA SpeedVac). (13) Add Hi-Di™ formamide (cold) 15 µl and keep at room temperature for 5 min.

835

Paragonimus

(14) Mix by pipetting ten times and transfer the contents of PCR tubes one by one to 96-well PCR reaction plate. Cover the PCR reaction plate with heat-resistant cover. (15) Heat the PCR plate at 96°C in thermocycler for 3 min then quickly remove and place on ice for 5 min. (16) Finally transfer the PCR reaction plate to the automatic DNA sequencer for capillary electrophoresis. Prior to that, remove the heat resistant cover and put 96-well plate septa on the plate and assemble the plate base. Load the PCR reaction plate in the sequencer. Note: The bench surface where PCR reactions are set up should always be wiped with alcohol prior to work to avoid contamination. Addition of trehalose (5% final concentration in reaction mix) makes it possible to store aliquots of master mixes at -20°C for direct addition to 96-well PCR plates and storage at -20°C for 1 month. Plugged tips should be used for all PCR reagents.

61.3 Conclusions and Future Perspectives Clearly, the application of PCR approaches and DNA sequencing of lung-flukes is proving to be an indispensable tool for diagnosis and unambiguous identification of Paragonimus species. It is possible to obtain sufficient DNA from a small number of eggs for diagnosis to the level of species, and further refinement and improvement of such methods should continue. Of course, eggs are not always obtained easily, even in cases of pulmonary paragonimiasis. Immunodiagnostic tests will therefore continue to be very important, especially for detection of ectopic paragonimiasis. Future interest in diagnosis of lung-fluke infection will also be sustained by the need to distinguish between paragonimiasis and tuberculosis.











References











1. Blair, D. et al. Paragonimiasis and the genus Paragonimus. Adv Parasitol., 42, 113, 1999. 2. Blair, D. et al. Paragonimiasis. In World Class Parasites: Volume 11, Food-Borne Parasitic Zoonoses. Murrell, K.D. and Fried, B. (Eds.). Springer, Berlin, Germany, 2007. 3. Miyazaki, I. An Illustrated Book of Helminthic Zoonoses. International Medical Foundation of Japan, Tokyo, 1991. 4. Yoshikawa, M. Immunopositivity for Trichinella spiralis presented in a case of Paragonimiasis westermani – suspected infection by ingestion of raw bear meat. J. Nara Med. Assoc., 56, 253, 2005. 5. Keiser, J. and Utzinger, J. Emerging foodborne trematodiasis. Emerg. Infect. Dis., 11, 1507, 2005. 6. Fried, B. et al. Food-borne intestinal trematodiases in humans. Parasitol. Res., 93, 159, 2004. 7. Toscano, C. et al. Paragonimiasis and tuberculosis, diagnostic confusion: a review of the literature. Trop. Dis. Bull. 92, R1, 1995. 8. Singh, T.N. et al. Pleuropulmonary paragonimiasis mimicking pulmonary tuberculosis – a report of three cases. Indian J. Med. Microbiol., 23, 131, 2005.



9. Narain, K. et al. Pulmonary paragonimiasis and smear-negative pulmonary tuberculosis: a diagnostic dilemma. Int. J. Tubercul. Lung Dis., 8, 621, 2004. 10. Mahajan, R.C. Paragonimiasis: an emerging public health problem in India. Indian J. Med. Res., 121, 716, 2005. 11. Sharma, D.C. Paragonimiasis causing diagnostic confusion with tuberculosis. Lancet Infect. Dis., 5, 538, 2005. 12. Singh, T.N. et al. Pulmonary paragonimiasis. Indian J. Chest Dis. Allied Sci., 46, 225, 2004. 13. Olson, P.D. et al. Phylogeny and classification of the Digenea (Platyhelminthes: Trematoda). Int. J. Parasitol., 33, 733, 2003. 14. Blair, D. et al. A molecular perspective on the genera Paragonimus Braun, Euparagonimus Chen and Pagumo­ gonimus Chen. J. Helminthol., 73, 295, 1999. 15. Blair, D. et al. Phylogenetic relationships among the Thai species of Paragonimus inferred from DNA sequences. In Proceedings, IX International Congress of Parasitology, Chiba, Japan, pp. 643–647. Tada, I. et al. (Eds.). Monduzzi Editore Sp. A., Bologna, Italy, 1998. 16. Binchai, S. et al. Molecular systematics of a new form of Paragonimus westermani discovered in Thailand. SE Asian J. Trop. Med. Pub. Health, 28, 92, 2007. 17. Sugiyama, H. et al. New form of Paragonimus westermani discovered in Thailand: morphological characteristics and host susceptibility. SE Asian J. Trop. Med. Pub. Health, 38, 2007. 18. Blair, D. et al. Geographical genetic structure within the human lung fluke, Paragonimus westermani, detected from DNA sequences. Parasitol. 115, 411, 1997. 19. Blair, D. et al. Molecular evidence for the synonymy of three species of Paragonimus, P. ohirai Miyazaki, 1939, P. iloktsuenensis Chen, 1940 and P. sadoensis Miyazaki et al., 1968. J. Helminthol., 71, 305, 1997. 20. Blair, D. et al. Paragonimus skrjabini Chen, 1959 (Digenea: Paragonimidae) and related species in eastern Asia: a combined molecular and morphological approach to identification and taxonomy. Syst. Parasitol., 60, 1, 2005. 21. Iwagami, M. et al. Identities of two Paragonimus species from Sri Lanka inferred from molecular sequences. J. Helminthol., 77, 239, 2003. 22. Iwagami, M. et al. Ancient divergence of Paragonimus westermani in Sri Lanka. Parasitol. Res., 102, 845, 2008. 23. Tandon, V. et al. Surface fine topography and PCR-based determination of metacercaria of Paragonimus sp. from edible crabs in Arunachal Pradesh, Northeast India. Parasitol. Res., 102, 21, 2007. 24. Thaenkham, U. and Waikagul, J. Molecular phylogenetic relationship of Paragonimus pseudoheterotremus. SE Asian J. Trop. Med. Pub. Health, 39, 217, 2008. 25. Nolan, M.J. and Cribb, T.H. The use and implications of ribosomal DNA sequencing for the discrimination of digenean species. Adv. Parasitol. 60, 101, 2005. 26. Owen, I.L. Parasitic zoonoses in Papua New Guinea. J. Helminthol., 79, 1, 2005. 27. Agatsuma, T. et al. Population genetic studies on isozymes in Paragonimus westermani from the Philippines. Jpn. J. Parasitol., 37, 195, 1988. 28. Agatsuma, T. et al. Genetic differentiation between Malaysian and other Asian Paragonimus westermani. Trop. Biomed. 10, 45, 1993. 29. Singh, S.T. et al. Possible discovery of Chinese lung fluke, Paragonimus skrjabini in Manipur, India. SE Asian J. Trop. Med. Pub. Health, 37, 53, 2006.

836 30. Zhou, B.J. et al. Sequence analyses of ITS2 and CO1 genes of Paragonimus proliferus obtained in Yunnan province, China and their similarities with those of P. hokuoensis. Parasitol. Res., 102, 1379, 2008. 31. Devi, K. et al. Pleuropulmonary paragonimiasis due to Paragonimus heterotremus: molecular diagnosis, prevalence of infection and clinicoradiological features in an endemic area of northeastern India. Trans. Roy. Soc. Trop. Med. Hyg., 101, 786, 2007. 32. Singh, T.S. et al. Morphological and molecular charac­ terizations of Paragonimus heterotremus, the causative agent of human paragonimiasis in India. SE Asian J. Trop. Med. Pub. Health, 38, 82, 2007. 33. Le, T.H. et al. Paragonimus heterotremus Chen and Hsia (1964), in Vietnam: a molecular identification and relation­ ships of isolates from different hosts and geographical origins. Acta Trop., 98, 25, 2006. 34. Miyazaki, I. and Fontan, R. Mature Paragonimus heterotremus found from a man in Laos. Jap. J. Parasitol., 19, 109, 1970. 35. Simarro, P.P. et al. Endemic human paragonimiasis in Equatorial Guinea: Detection of the existence of endemic human paragonimiasis in Equatorial Guinea as a result of an integrated sanitary programme. Trop. Geog. Med., 43, 326, 1991. 36. Oelerich, S. and Volkmer, K.J. Antikorper- und Immunog­ lobulinbestimmungen bei Patienten mit afrikanischer Para­ gonimiasis. Tropenmed. Parasitol., 27, 44, 1976. 37. Kum, P.N. and Nchinda, T.C. Pulmonary paragonimiasis in Cameroon. Trans. Roy. Soc. Trop. Med. Hyg., 76, 768, 1982. 38. Voelker, J. et al. On the epidemiology of paragonimiasis in man and in animals in Nigeria, West Africa. Vet. Med. Rev., 1/2, 161, 1975. 39. Nozais, J.P. et al. Paragonimiasis in Africa. A recent focus in Ivory Coast. Bull. Soc. Pathol. Exot., 73, 155, 1980. 40. Sachs, R. et al. Le Paragonimus uterobilateralis comme cause de trois cas de paragonimose humaine au Gabon. Tropenmed. Parasitol., 34, 105, 1983. 41. Voelker, J. and Nwokolo, C. Human paragonimiasis in eastern Nigeria caused by Paragonimus uterobilateralis. Z. Tropenmed. Parasit., 24, 323, 1973. 42. Onuigbo, W.I. and Nwako, F.A. Discovery of adult parasites of Paragonimus uterobilateralis in human tissue in Nigeria. Tropenmed. Parasitol., 25, 433, 1974. 43. Udonsi, J.K. Clinical field trials of praziquantel in pulmonary paragonimiasis due to Paragonimus uterobilateralis in endemic populations of the Igwun Basin, Nigeria. Tropenmed. Parasitol., 40, 65, 1989. 44. Sachs, R. and Voelker, J. Human paragonimiasis caused by Paragonimus uterobilateralis in Liberia and Guinea, West Africa. Tropenmed. Parasitol., 33, 15, 1982. 45. Monson, M.H. et al. Successful treatment with praziquantel of six patients infected with the African lung fluke, Paragonimus uterobilateralis. Am. J. Trop. Med. Hyg., 32, 371, 1983. 46. Marty, A.M. and Neafie, R.C. Paragonimiasis. In Pathology of Infectious Diseases: Volume I—Helminthiases, pp 49–67. Meyers, W.M. et al. (Eds.). Armed Forces Institute of Pathology and American Registry of Pathology, Washington DC, 2000. 47. Procop, G.W. et al. North American paragonimiasis—A case report. Acta Cytol., 44, 75, 2000. 48. DeFrain, M. and Hooker, R. North American paragonimiasis— Case report of a severe clinical infection. Chest, 121, 1368, 2002. 49. Zarrin-Kameh, N. et al. Pulmonary paragonimiasis diagnosed by fine needle aspiration. J. Clin. Microbiol., 46, 2137, 2008. 50. Joshi, S. and Rao, M.K.K. Paragonimiasis in Manipur. Med. J. Armed Forces India, 50, 132, 1994.

Molecular Detection of Foodborne Pathogens 51. Yokogawa, S. et al. Paragonimus and paragonimiasis. Exper. Parasitol., 10, 81 &139, 1960. 52. Yokogawa, M. Paragonimus and paragonimiasis. Adv. Parasitol., 3, 99, 1965. 53. Zhong, H.L. et al. Recent progress in studies of Paragonimus and paragonimiasis control in China. Chinese Med. J., 94, 483, 1981. 54. Quicho, L.P. et al. Humoral immune response of cats to Paragonimus infection. SE Asian J. Trop. Med. Pub. Health, 12, 364, 1981. 55. Waikagul, J. Serodiagnosis of paragonimiasis by enzymelinked immunosorbent assay and immunoelectrophoresis. SE Asian J. Trop. Med. Pub. Health, 20, 243, 1989. 56. Maleewong, W. et al. Comparison of adult somatic and excretory-secretory antigens in enzyme-linked immunosorbent assay for serodiagnosis of human infection with Paragonimus heterotremus. Trans. Roy. Soc. Trop. Med. Hyg., 84, 840, 1990. 57. Wongkham, C. et al. Partially purified antigens of Paragonimus heterotremus for serodiagnosis of human paragonimiasis. SE Asian J. Trop. Med. Pub. Health, 25, 176, 1994. 58. Maleewong, W. et al. Indirect haemagglutination test using monoclonal antibody-affinity purified antigens for diagnosis of human paragonimiasis due to Paragonimus heterotremus. Trop. Med. Internat. Health, 3, 57, 1998. 59. Kim, T.Y. et al. Recombinant Paragonimus westermani yolk ferritin is a useful serodiagnostic antigen. J. Infect. Dis., 185, 1373, 2001. 60. Cho, S.Y. et al. Antibody changes in paragonimiasis patients after praziquantel treatment as observed by ELISA and immunoblot. Korean J. Parasitol., 27, 15, 1989. 61. Zhang, Z.H. et al. Characterization of stage-specific antigens of Paragonimus westermani. Am. J. Trop. Med. Hyg., 44, 108, 1991. 62. Slemenda, S.B. et al. Diagnosis of paragonimiasis by immunoblot. Am. J. Trop. Med. Hyg., 39, 469, 1988. 63. Sugiyama, H. et al. Characterization and localization of Paragonimus westermani antigen stimulating antibody formation in both the infected cat and rat. J. Parasitol., 73, 363, 1987. 64. Itoh, M. and Sato, S. An antigenic component for the serodiagnosis of paragonimiasis miyazakii. Jpn. J. Parasitol., 37, 347, 1988. 65. Kim, S.H. et al. Immunoblot observation of antigenic protein fractions in Paragonimus westermani reacting with human sera. Korean J. Parasitol., 26, 239, 1988. 66. Dekumyoy, P. et al. Human lung fluke Paragonimus heterotremus: differential diagnosis between Paragonimus heterotremus and Paragonimus westermani infections by EITB. Trop. Med. Internat. Health, 3, 52, 1998. 67. Maleewong, W. et al. Detection of Paragonimus heterotremus in experimentally infected cat feces by antigen capture-ELISA and by DNA hybridization. J. Parasitol., 83, 1075, 1997. 68. Chang, Z.S. et al. Gene sequences for identification of Paragonimus eggs from a human case. Chinese J. Parasitol. Parasit. Dis., 18, 213, 2000. 69. Sugiyama, H. et al. Polymerase chain reaction (PCR)-based molecular discrimination between Paragonimus westermani and P. miyazakii at the metacercarial stage. Mol. Cell. Probes, 16, 231, 2002. 70. Sugiyama, H. et al. Molecular discrimination between individual metacercariae of Paragonimus heterotremus and P. westermani occurring in Thailand. SE Asian J. Trop. Med. Pub. Health, 36, 102, 2005. 71. Gyoten, J. Infection by Paragonimus miyazakii cercariae of their crab hosts, Geothelphusa dehaani, by percutaneous penetration. J. Parasitol., 86, 1342, 2000.

Paragonimus 72. Miyazaki, I. Lung flukes in the world – Morphology and life history. In A Symposium on Epidemiology of Parasitic Diseases, pp 101–135. Sasa, M. (Ed.). International Medical Foundation of Japan, Tokyo, 1974. 73. Le, T.H. et al. Human fascioliasis and the presence of hybrid/ introgressed forms of Fasciola hepatica and Fasciola gigantica in Vietnam. Int. J. Parasitol., 38, 725, 2008. 74. Castilla, E.A. et al. Cavitary mass lesion and recurrent pneumothoraces due to Paragonimus kellicotti infection – North American paragonimiasis. Am. J. Surg. Pathol., 27, 1157, 2003. 75. Jun, S.Y. Paragonimiasis of the breast. Report of a case diagnosed by fine needle aspiration. Acta Cytol., 47, 685, 2003. 76. Hiratsuka, T. et al. Paragonimiasis westermani with repeated pleural effusion and diagnosed by detecting parasite eggs in pleural effusion. Nihon Kokyuki Gakkai Zasshi, 45, 49, 2007. 77. Kim, J.Y. et al. Laparoscopic excision of intra-abdominal paragonimiasis. Surg. Laparosc. Endosc. Percut. Technol., 17, 556, 2007. 78. Schuster, H. et al. Otitis media and a neck lump – Current diagnostic challenges for Paragonimus-like trematode infections. J. Infect., 54, e103, 2006. 79. Intapan, P.M. et al. Detection of Paragonimus heterotremus eggs in experimentally infected cats by a polymerase chain reaction-based method. J. Parasitol., 91, 195, 2005.

837 80. Sugiyama, H. et al. Application of multiplex PCR for species discrimination using individual metacercariae of Paragonimus occurring in Thailand. SE Asian J. Trop. Med. Pub. Health, 37, 48, 2006. 81. Zhang, Z. et al. Diagnosis of active Paragonimus westermani infections with a monoclonal antibody-based antigen detection assay. Am. J. Trop. Med. Hyg., 49, 329, 1993. 82. Zhang, Z. et al. Antigen detection assay to monitor the ­efficacy of praziquantel for treatment of Paragonimus westermani infections. Trans. Roy. Soc. Trop. Med. Hyg., 90, 43, 1996. 83. Kim, T.Y. et al. Recombinant Paragonimus westermani yolk ferritin is a useful serodiagnostic antigen. J. Infect. Dis., 185, 1373, 2002. 84. Lee, C.H. et al. Serial CT findings of Paragonimus infested dogs and the micro-CT findings of the worm cysts. Korean J. Radiol., 8, 372, 2007. 85. Maleewong, W. et al. Excretory-secretory antigenic components of Paragonimus heterotremus recognized by infected human sera. J. Clin. Microbiol., 30, 2077, 1992. 86. Yun, D.H. et al. Structural and immunological characteristics of a 28-kilodalton cruzipain-like cysteine protease of Paragonimus westermani expressed in the definitive host stage. Clin. Diag. Lab. Immunol., 7, 932, 2000.

62 Taenia

Akira Ito, Minoru Nakao, Yasuhito Sako, Kazuhiro Nakaya, and Tetsuya Yanagida Asahikawa Medical College

Munehiro Okamoto Tottori University

Contents 62.1. Introduction..................................................................................................................................................................... 839 62.1.1. Classification, Biology, and Pathogenesis of Foodborne Taenia....................................................................... 840 62.1.2. Present Situation of Taeniasis in Human and Animal Hosts............................................................................ 840 62.1.3. Current Techniques for Diagnosis of Taenia..................................................................................................... 840 62.1.3.1. Molecular Approaches...................................................................................................................... 841 62.1.3.2. Immunological Approaches.............................................................................................................. 842 62.1.3.3. Relative Performance of Molecular and Immunological Techniques.............................................. 842 62.2. Methods........................................................................................................................................................................... 843 62.2.1. Reagents, Supplies, and Equipment.................................................................................................................. 843 62.2.2. Sample Collection and Preparation................................................................................................................... 843 62.2.3. Detection Procedures........................................................................................................................................ 845 62.2.3.1. Multiplex PCR.................................................................................................................................. 845 62.2.3.2. Sequencing of the PCR Product....................................................................................................... 846 62.3. Conclusions and Future Perspectives.............................................................................................................................. 847 Acknowledgments...................................................................................................................................................................... 847 References.................................................................................................................................................................................. 848

62.1  Introduction Cestodes constitute one of the major groups in phylum Platyhelminthes.1 All cestode species infective to humans gain entry via oral ingestion of either eggs, which develop into the larval stage (metacestodes) in parenteral tissues and cause metacestodiases, or metacestodes, which develop into adult cestodes or tapeworms in the intestinal lumen and cause cestodiases. All cestode species but Hymenolepis nana require one or more intermediate hosts for completion of the life cycle.2,3 Pathologically, metacestodiases caused by ingested eggs are more serious and life-threatening zoonotic diseases, since the completion of the life cycle requires humans as well as other animals as the obligative intermediate or definitive host. The major metacestodiases in humans are cysticercosis and echinococcosis whose route of infection is oral ingestion of eggs. Being a foodborne disease, human sparganosis results from ingestion of uncooked or undercooked snakes or other animals harboring sparganum or spargana. Human is the paratenic host in which the parasite usually does not develop into adult tapeworms. Echinococcosis is generally not regarded a foodborne disease, however, its eggs expelled from dogs or foxes can be accidentally ingested by humans,

causing chronic, but lethal hepatic diseases.4 By contrast, human cysticercosis (and cestodiasis) is a foodborne disease as humans are infected either by ingestion of food contaminated with feces containing eggs (or by proglottids carried into the stomach by vomiting), or by eating undercooked or uncooked meat (pork and beef) harboring metacestodes of T. solium, T. saginata or T. asiatica. Tapeworm infections by Taenia spp, Diphyllobothrium spp., Diprogonoporus spp., Mesocestoides spp., Spirometra spp., and others are through consumption of uncooked or undercooked foods or wild animals contaminated with metacestodes of these cestode species.5 Among them, Taenia spp. are the major foodborne pathogens, since metacestode stage of these species are in the major food or meat products for human consumptions. Taenia solium is unique and a most serious pathogen, since it causes neurocysticercosis (NCC) due to the accidental ingestion of eggs. However, the eggs are released from humans exclusively who ate undercooked or uncooked pork harboring metacestodes of T. solium. In this chapter, we focus on three human Taenia species, T. solium, T. saginata and T. asiatica, the third species recently described from Asia.6 As NCC is one of the most important parasitic diseases,7,8 it will be also discussed. 839

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Molecular Detection of Foodborne Pathogens

62.1.1 Classification, Biology, and Pathogenesis of Foodborne Taenia

62.1.2 Present Situation of Taeniasis in Human and Animal Hosts

Human taeniasis caused by the so-called beef tapeworm, Taenia saginata and pork tapeworm, T. solium has a cosmopolitan distribution. The geographic distributions of these two species are somewhat different, as T. saginata is common where people eat raw or undercooked beef, whereas T. solium is common where they eat raw or undercooked pork. Larval stage, metacestode or cysticercus/cysticerci of these two species develop into adult tapeworm(s) in the intestine of humans, the only definitive hosts. The most important intermediate host of T. saginata is cattle, but buffaloes, giraffes, llamas, and reindeer may serve as the intermediate hosts, whereas that of T. solium is mainly pigs. However, dogs9, and humans are also highly susceptible hosts when these hosts have chances to ingest viable eggs orally.6,9 Therefore, humans are the only host in which T. solium develop into metacestodes from eggs as the intermediate host, and adult tapeworms from metacestodes as the definitive host. There is no confirmed evidence of human cysticercosis due to metacestodes of T. saginata.6,10 From a public health perspective, T. solium is very important, since humans may be infected with metacestodes after ingestion of eggs causing cysticercosis, especially NCC.7 The life cycle of these tapeworms is highly dependent on the consumption of contaminated food (meat) by humans. Therefore, tapeworm infections are eradicable if only ­parasite-free beef or pork or well cooked meat are consumed.11 It is simple to cut off the life cycle: not to eat uncooked or undercooked meat contaminated with metacestodes infective to humans. Taenia infections, especially T. solium in pigs are uncommon in developed countries nowadays, where pigs are raised under industrial environment which effectively blocks direct contact of pigs to human feces contaminated with eggs of this parasite. However, pigs, and dogs and other domestic animals are roaming with free access to human feces in developing countries. So, dogs as well as well-known pigs may harbor metacestodes in developing countries where T. solium is endemic.9,11,12 The situation with T. saginata is different. In developed countries, meat inspection has in general been well established for production of the safe beef. Cattle are grown in the pasture where strangers cannot access. Nonetheless, even under such conditions, some sporadic cases of cattle contaminated with metacestodes of T. saginata have been reported in developed countries such as USA or Canada. On the other hand, cattle in developing countries are grown in the open fields that may easily be contaminated by tapeworm carriers. A third species, T. asiatica, has been described from people in Asia and the Pacific. Morphologically, it is impossible to differentiate T. asiatica from T. saginata.6,14–21 There is a serious debate on its taxonomic independence from the species or strain of T. saginata.6,17,18,22 The tapeworm carriers are mainly the local people who have customs to eat uncooked or undercooked viscera of pigs.

Human taeniasis and cysticercosis of T. saginata and T. solium are well documented. T. saginata is cosmopolitan, because it is found in a majority of countries (except Hindi in India and Nepal) where beef forms a part of staple diet.23–25 T. solium is also cosmopolitan but it does not generally distribute in countries (e.g., the Middle East) where pork is not consumed.26 However, outbreaks of cysticercosis of T. solium have been reported in Saudi Arabia27 and in Kuwait.7,28,29 T. asiatica has been reported from Taiwan, Korea, China, Vietnam, Philippines, Indonesia, Malaysia, and Thailand.6,30,31 T. asiatica is distributed in Burma with no doubt.31 In Asia and the Pacific, there was a long standing puzzle “Why do local people eating no beef but pork harbor T. saginata?.” Now, it is clear that it is due to the infection of the third species, T. asiatica. Therefore, we have to re-evaluate T. saginata from Asia and the Pacific, if they are real T. saginata or T. asiatica or both, since three species may sympatrically be distributed in several countries.6,30,31 In other areas in the world, two species, T. saginata and T. solium are distributed through eating meat. However, many tapeworm carriers treated in Americas and Europe or even in Africa may have history to visit Asia and the Pacific or immigrants from Asia. Therefore, we have to re-examine these cases if the worms are real T. saginata or T. asiatica with questionnaire of history of eating under cooked viscera of pigs in Asia and the Pacific. Given the cost for inspection of domestic animals at slaughterhouses and the cost of reagents, serology to detect pigs infected with T. solium has not been widely applied. In addition, the existing serological methods for detection of individual cattle infected with metacestodes of T. saginata are not sensitive enough to be a practical tool. There is a good review on this topic with back ground socio-economic problems in developing countries.32 Obviously, future development of more sensitive and specific methodologies is necessary in order to determine how many domestic animals are contaminated with these metacestodes, although basic tools are available for detection in individuals infected with taeniasis related to “WAFP.” The term “WAFP” was proposed by Ito33 to describe “any kind of infectious disease, either emergent or re-emergent, may be caused through contaminated water, contaminated air, contaminated food or contaminated people.” The wave of globalization in human activities including enormous increases in international trading of foods and live stocks, in the number of travelers, immigrants, and refugees, poses new risks for many infectious diseases including taeniasis to emerge and/or re-emerge worldwide.

62.1.3 Current Techniques for Diagnosis of Taenia In this section, we review current molecular approaches for detection or identification of human taeniid cestodes, T. solium, T. saginata and T. asiatica (Asian Taenia), as well as immunological approaches for detection of cysticerci or adult

841

Taenia

(a)

(b)

500 µm

Figure 62.1  Scoleces of T. solium (a) and T. asiatica (b) expelled from one woman in Thailand. Two T. solium and one T. asiatica were identified by mitochondrial DNA analysis. (From Ananthapuruti, M.T. et al., Emerg. Infect. Dis., 13, 1413, 2007.)

tapeworms in infected humans and animals. Morphological identification of the species is still essential. Figure 62.1 illustrates two types of Taenia scoleces collected from a patient in Thailand.31 One (A) was expected to be T. solium with armed hooks, whereas the other (B) was to be T. saginata, since there was no report to indicate the distribution of T. asiatica in Thailand, at that time. However, molecular analyses revealed that the latter was not T. saginata but T. asiatica, and three species, T. solium, T. saginata and T. asiatica occurred sympatrically.31 62.1.3.1  Molecular Approaches (i) Molecular identification of taeniid proglottids expelled from humans: Molecular diagnostic techniques have played a vital role in the confirmation of the third human taeniid species, Asian Taenia.14 Details in the history of findings of the Asian Taenia have just been reviewed by Flisser and others.34 Zarlenga and others35 and Bowles and McManus22 stressed the molecular difference between the classic T. saginata and Taiwan Taenia was too small and should be considered to be intraspecific variations, even though one of the authors, Fan, and others36 wanted to describe it as a new species. Later, it was designated as a new independent species, T. asiatica by other group.14–17,20,34,37 (ii) Two genotypes of T. solium in the world: A Japanese group uncovered that T. solium worldwide could be differentiated into two, Asian and American/African genotypes.38–40 Based on the sequence data of the full-length mitochondrial DNA of T. solium,39 a multiplex PCR was developed by the same group,40–42 which enabled differentiation of the two genotypes of T. solium in the world: the Asian type, and the American/African type. It has been conceived that the clinical manifestation of cysticercosis in Asia and the Pacific often includes NCC and subcutaneous cysticercosis (SCC),6,44–46 whereas it mainly includes NCC without SCC in America and Africa with some rare exceptions. However, NCC without SCC in Asia, and NCC with SCC in Africa, are also often observed and reported. Therefore, the earlier concept might be somewhat biased arising from a limited number of

cases.7,46 Although there is no crucial evidence of the correlation of genotypes and clinical manifestations,7,46 such genotyping is expected to be informative for tracking back where NCC patients were exposed to T. solium eggs. Most recent studies on mitochondrial DNA of T. solium have revealed subtypes in these two genotypes.47,48 Sudewi and others48 analyzed the mtDNA patterns using ­paraffin-embedded cysts resected from a tongue or from subcutaneous nodule of two Balinese men and found some crucial difference from other isolates in Asia. Such molecular confirmation using histopathological specimens 7,43,49–51 or cerebrospinal fluid (CSF)52 may give us useful information where the patients had the risk of infections. (iii) Histopathological specimens for molecular identification: NCC is serious and histopathological examination of the causative tumor-like cysts in the brain is not always available for identification. Immunological diagnosis for NCC is sometimes ineffective, as detection of antibodies is not always positive53,54 especially when the patient has a solitary cyst. In addition, if the patient suffers from certain immunosuppressed conditions, antibody response may be poor or none. Some other zoonotic taeniid metacestodes may also accidentally infect humans. In cases that histopathological specimens are available, the characteristic Taenia hooklets may not always be revealed. Furthermore, when severe combined immunodeficient (scid) mice were experimentally infected with in vitro hatched oncospheres of T. solium from Asia, the resulting metacestodes showed high variation of hooklets from zero to 34 (double lings) (Ito et al. unpublished).9 Therefore, observation of hooklets does not always provide a conclusive means to identify T. solium, keeping in mind other species which might infect humans as very rare cases.7,49 Thus, when we have chances to examine histopathological specimens from patients, it is strongly recommended to analyze the mtDNA for species and genotype identification of T. solium. (iv) Racemose cysticercosis: The mechanism of atypical, acephalic, racemose form of NCC is not totally understood. However, such cases might be more common in immunocompromized patients.7,50,51 Therefore, molecular tools should be applied for identification of the racemose form of NCC.55 (v) Molecular tools for epidemiology to prevent NCC: Multiplex PCR is valuable for differentiation not only of the two genotypes of T. solium but also two other human taeniid species, T. saginata and T. asiatica, which may sympatrically be occurring in some areas in Asia and the Pacific.29–31,41,56 Molecular tools are necessary for the identification and real-time analysis of the worm carriers and ­prevention of accidental NCC in the endemic areas. Although copro-ELISA tests have been used for screening of taeniasis,57 molecular identification is essential for assessing treatment efficacy. While multiplex PCR can also be utilized for detection of eggs in feces,41,58 it is not applicable in endemic but remote areas where the electric power is unreliable. Therefore, much simpler tool, loop-­mediated ­isothermal amplification (LAMP)59 for differentiation of

842

three human Taenia sympatrically occurring in Asia and the Pacific has most recently been established by Nkouawa et al.60 (vi) Other zoonotic taeniid infections: The majority of Taenia species causing cysticercosis in humans is known to be T. solium. However, the existence of many other zoonotic taeniid cestode species including T. taeniaeforimis, T. pisiformis, T. hydatigena, T. serialis, and T. multiceps may add complexity in the diagnosis.7,42 Many cox1 gene sequences from not only taeniid cestodes but also various organisms have been deposited in the database. When we use universal primer sets such as JB3/JP4.522 or pr-a/pr-b,61 it may allow us to determine the causative organisms. Further studies to establish additional molecular probes for identification of other zoonotic cestodes species may be invaluable when we encounter such unexpected species.7,62 62.1.3.2  Immunological Approaches (i) T. solium cysticercosis in humans, pigs, and dogs or other wild animals: For the screening of cysticercosis in humans and domestic animals, serology offers a useful tool to identify the individuals infected with cysticerci of these species. Both neuroimaging and serology are highly informative for diagnosis.7,8 The most reliable and practical serological tools have been developed for detection of human cysticercosis. Glycoproteins (GPs) highly specific to T. solium cysticercosis have been utilized. Recombinant antigens and synthetic peptides have also been produced and applied for routine serodiagnosis.63–73 Serological detection of pigs and dogs infected with T. solium cysticerci has been reported using purified GPs and recombinant chimeric antigens which are highly useful for detection of cysticercosis patients.13,70–75 So far, these antigens prepared by research group at Asahikawa Medical College were applicable not only for immunoblots but also for ELISA with almost the same sensitivity and specificity. The GPs at approximately PI at 8.0–9.0 purified by isoelectric focusing were highly specific and useful for both immunoblot and ELISA.66,72–75 The GPs purified by lentil lectin-affinity chromatography were recommended only for immunoblots,64,76 as they tended to cross-react with nonspecific components in ELISA. Nonetheless, the immunoblot using GPs purified by lentil lectin-affinity chromatography was the gold standard from 1990 onwards. The synthetic peptides prepared by the same group seemed to provide excellent resolution for both immunoblot and ELISA.71,77,78 Alternative approach to detect antibodies in humans and domestic animals is to apply crossreactive antigens from metacestodes of T. crassiceps79,80 or T. saginata.81 As such candidate antigens are hydrophobic ligand binding proteins (HLBP) with high homology among taeniid cestodes, molecular basis on such heterologous antigens should be well documented for publication.82 Another approach to detect infected humans or animals is the detection of circulating antigens of T. solium.83–85 So far all tools for detection of circulating antigens are based on cross-reactions to monoclonal antibodies to some components of T. saginata.86,87 However, there is no molecular

Molecular Detection of Foodborne Pathogens

information on the antigens captured by the monoclonal antibodies. There is no comparison of immunological tools to detect circulating antigens using monoclonal antibody to T. saginata and to detect antibodies using GPs or recombinant antigens of T. solium.88 (ii) NCC in humans: NCC is one of the most serious parasitic zoonoses threatening human life worldwide,63 and many challenges remain to establish sensitive and reliable immunodiagnostic techniques for this disease. There is a consensus among many researchers that a combination of serology and neuroimaging is essential for correct diagnosis of NCC.7,8,12,89,90 Although molecular confirmation provides crucial, reliable, and direct evidence, it is not always easy to obtain such parasite specimens from NCC cases. So, molecular identification is an option when specimens from patients are available. There are two strategies for NCC immunodiagnosis: using serum samples and CSF. Although many researchers previously stressed and recommended the use of CSF for confirmative immunodiagnosis, recent comparative studies using serum and CSF samples from the same patients showed negligible difference between these samples when recombinant antigens was applied.7,8 Advantage of using CSF for diagnosis for NCC may still have relevance to the detection of T. solium DNA.52 (iii) T. saginata cysticercosis (bovine cysticercosis) in cattle: Harrison et al.86 and Brandt et al.87 attempted to detect circulating antigens of T. saginata in cattle. The sensitivity appeared low, which might be due to the small number of metacestodes developed in experimentally infected cattle. Research groups in Europe or America or Canada might have problems in obtaining large quantity of fresh viable eggs for such experimental infections. It may be better carried out in endemic areas (e.g., Asia) where fresh T. saginata or T. asiatica worms and eggs are readily accessible.25,30,31 (iv) T. asisatica cysticercosis in pigs: There is no report as to how to detect T. asiatica in infected pigs. Experimental infections were carried out by Fan and others,36,91,92 whose main interest was to confirm development of metacestodes in the viscera of various domestic animal species but not to detect antibodies. Such experimental infections are also necessary in order to establish new tools how to detect infected animals in endemic areas. HLBP shared among taeniid cestode species may be alternative candidate antigens for detection of antibodies to T. asiatica in pigs. (v) Usefulness of immunodeficient mice as alternative intermediate hosts: Considering the challenges we face to obtaining fully developed metacestodes of human Taenia spp.93 (Figure 62.2), immunodeficient mice may represent a highly useful tool for generation of fully matured metacestodes for diagnostic and other applications.94,95 62.1.3.3 Relative Performance of Molecular and Immunological Techniques The critical differences between molecular and immunological approaches are that the former provides direct evidence

843

Taenia

(a)

(b)

(c)

Figure 62.2  Cysticerci of T. saginata (a), T. asiatica (b) and T. solium (c) fully developed in NOD/Shi-scid mouse (Bar = 5 mm). (Modified from Nakaya, K. et al. Parasitol. Int., 55, S91, 2006.). Those of T. saginata were exclusively confirmed to be infective to human volunteers (unpublished data).

while the latter provides indirect evidence. Immunological tools are based on how to detect specific antibodies against specific antigens or to detect specific antigens, either in the blood stream or in feces, using monoclonal or polyclonal antibodies to the candidate antigens. Therefore, it is impossible to differentiate T. solium, T. saginata, and T. asciatica immunologically without any crucial molecular evidence on the species-specific components based on careful studies using reliable materials from these three species. As humans are the only intermediate host of T. solium, if the antigen candidate is not really species specific, it still has value for detecting cysticercosis patients, since two other taeniid species do not infect humans through oral ingestion of eggs.6 So, if such immunological tools confirm the presence of metacestodes of any Taenia species in humans, it almost automatically indicates metacestode(s) of T. solium. Antigens from other species that do not infect humans except very few accidental cases may also be useful for detection of cysticercosis of T. solium. However, it is different in domestic animals, which may also be infected with metacestodes of other taeniid species.7,8 By contrast, molecular tools provide species-specific DNA information based on comparative studies of DNA sequences of all species deposited in GenBank. Even a SNP (single nuclear point) may be useful for multiplex PCR or other tools discussed above. So far, all molecular approaches to identify human Taenia infections are based on mitochondrial DNA analyses. However, microsatellite DNA studies96–98 are also expected to be informative for future studies. Combination of mtDNA information and nuclear DNA information including microsatellite DNA and other genes is essential in dealing with the taxonomic problem on Asian Taenia as if it is very closely related genetically to species or

strain of T. saginata. If mtDNA and nuclear DNA analysis of an Asian Taenia specimen reveals different origins, it is likely that this parasite may be hybrids between the Asian Taenia and T. saginata.21

62.2  Methods In order to identify taeniid species, various molecular approaches have been developed (e.g., DNA hybridization, PCR-RFLP, PCR-SSCP, RAPID-PCR, sequencing of DNA, etc.). Molecular probes for identification of species or genotypes of human Taenia summarized in Table 62.1. Each of these techniques has its own advantages and disadvantages. While DNA hybridization, PCR-RFLP, and PCR-SSCP are relatively time-consuming procedures, PCR using speciesspecific primers provides rapid and sensitive resolution.42

62.2.1 Reagents, Supplies, and Equipment General reagents, supplies, and equipment that are required for the molecular diagnosis of taeniid cestodes are listed in Table 62.2.

62.2.2 Sample Collection and Preparation Three scenarios are expected in the detection of human tapeworm carriers. The first is identification of individual cases who admit hospitals after some symptoms are recognized by the carriers themselves.6 The most reliable symptom is history of active expulsion of proglottids of T. saginata or T. asiatica. However, it is not so clear in T. solium carriers. In this case, proglottids expelled from the carrier should be submitted for morphological and molecular identification. Fecal

844

Molecular Detection of Foodborne Pathogens

Table 62.1 Molecular Probes for Identification of Human Taeniid Species Hybridization Southern, dot

PCR PCR-RFLP

Target DNA Unknown Unknown Noncoding DNA fragments Unknown: cox1 Target DNA HDP2 (170 bp) ITS1 (760 bp) ITS2 (Tsag: 650 bp) (Tsol: 600 bp ) cox1(521 bp)

PCR-SSCP

ND1 (530 bp) cox1 (444 bp)

RAPD SCAR

Unknown Unknown SCAR marker A03 (864 bp) SCAR marker AD14 (463 bp) SCAR marker AD16 (502 bp) SCAR marker D05 (545 bp) SCAR marker S07 (484 bp) SCAR marker G14 (377 bp) SCAR marker K02 (531 bp)

BESS T-base

cox1 (469 bp) cob (253 bp)

Specifc PCR

HDP1 (ladder)

Multiplex PCR

HDP2 ( Tsag:170 bp, 600 bp) (Tsol: 170 bp) cox1 (Tsag: 827 bp) (Tasi: 269 bp) (TsolAm/Af: 720 bp) (TsolAsia: 984 bp)

PCR/sequencing

28SrDNA D1 ( 281 bp) cox1 (444 bp) 12S rDNA (311 bp)

Probe pTS10 Total genomic DNA HDP1, HDP2 pTsol-9, pTsag-16

Differential Diagnosis T. solium specific T. solium, T. saginata T. solium, T. saginata T. solium, T. saginata

Primers PTs7S35F2: CTTCTCAATTCTAGTCGCTGTGGT PTs7S35R1: GGACGAAGAATGGAGTTGAAGGT BD1: GTCGTAACAAGGTTTCCGTA 4S: TCTAGATGCGTTCGAAG/ATGTCGATG NC6: ATCGACATCTTGAACGCACATTGC NC2: TTAGTTTCTTTTCCTCCGCT Forward primer: TGGGTTAGATGTTAAGACGGC Reverse primer: AAAACACCCACTAAAAGCAGA JB11: AGATTCGTAAGGGGCCTAATA JB12: ACCACTAACTAATTCACTTTC JB3: TTTTTTGGGCATCCTGAGGTTTAT JB4.5: TAAAGAAAGAACATAATGAAAATG Primers from Operon Technologies Primers from Operon Technologies A03F: AGTCAGCCACCTGTACACATG A03R: AGTCAGCCACGCAAAGGGTAG AD14F: GAACGAGGGTGAATCACATAGC AD14R: GAACGAGGGTCGATGTATAAAT AD16F: AACGGGCGTCCTAGGAGATGGC AD16R: AACGGGCGTCAGTAGTGAAC D05F: TGAGCGGACACCGAAAGCTATTGC D05R: TGAGCGGACAAGTAGTGAAGTG S07F: TCCGATGCTGAGCATGGGTAG S07R: TCCGATGCTGGTCAAAGTGGAAAG G14F: GGATGAGACCGTACTAGCATG G14R: GGATGAGACCCTTGTAACAAATG K02F: GTCTCCGCAACTGTAATGTATC K02R: GTCTCCGCAATAATAATCTTTC P1: ATATTTACTTTAGATCATAAGCG P2: TAAAATTAATAGAACTAAAAAT P3: TTATGAGATTGTCAAAAGATTCTT P4: TATAGATGTCAAAACAGTAGCAGCCC PTs4F1: GCAGTGTGCTGAAGATGAATA PTs4R1: GAATTTGGCTCTCACTGAATG PTs7S35F1: CAGTGGCATAGCAGAGGAGGAA PTs7S35F2: CTTCTCAATTCTAGTCGCTGTGGT PTs7S35R1: GGACGAAGAATGGAGTTGAAGGT TsagMpF: TTGATTCCTTCGATGGCTTTTCTTTTG TasiMpF: ACGGTTGGATTAGATGTTAAGACTA TsolAmMpF: GGTAGATTTTTTAATGTTTTCTTTA TsolAsiMpF: TTGTTATAAATTTTTGATTACTAAC TaeniaMpR: GACATAACATAATGAAAATG JB10: GATTACCCGCTGAACTTAAGCATAT JB9: CCTGCATCACAAACACCCCGACTC JB3: TTTTTTGGGCATCCTGAGGTTTAT JP4.5: TAAAGAAAGAACATAATGAAAATG 60.for.: TTAAGATATATGTGGTACAGGATTAGATACCC 375.rev.: AACCGAGGGTGACGGGCGGTGTGTACC

Differential diagnosis T. solium, T. sagniata

Reference 103 104 105 106 Reference 107

T. solium, T. sagniata T. solium, T. sagniata T. solium, T. sagniata

58

Taenia

108

Variation of T. solium T. solium, T. sagniata T. solium specific

109 110 110*

T. solium specific T. solium specific T. solium specific T. solium specific T. saginata spcific T. saginata spcific T. saginata, T. asiatica two genotyps of T. solium

102

T. saginata specific

111

T. solium, T. sagniata

111

T. saginata, T. asiatica two genotyps of T. solium

41*

Universal

22

Universal Taenia, Echinococcus

112

(Continued)

845

Taenia

Table 62.1  (Continued) PCR

Target DNA

Primers

cox1 (444 bp) cox1 (875 bp) ND4, ATP6, ND2 (1628 bp) cox1 (complete > 1620 bp) cob (complete  > 1068 bp)

pr-a: TGGTTTTTTGTGCATCCTGAGGTTTA pr-b: AGAAAGAACGTAATGAAAATGAGCAAC F/CO1: TTGAATTTGCCACGTTTGAATGC R/CO1: GAACCTAACGACATAACATAATGA F/ND4: TGGAGTTAGATGGTAAGCGTTGAT R/ND2: CAGGAAACTTCATAACAACACTTA Cox1/F: GTTATGTTAGACTAGATGTTTTCA Cox1/R: TCCACTAAGCATAATGCAAAAGGC Cytb/F: ATAAACTGATAGATTGTGGTTC Cytb/R: CATATGACTGTCTAATGAAGAAAA

Differential Diagnosis

Reference

Universal

61

Taenia, Echinococcus

113

Taenia T. saginata, T. asiatica two genotyps of T. solium

39

* Each primer was not named in the original paper.

Table 62.2 General Reagents, Supplies, and Equipment Required for Molecular Diagnosis Reagents Taq polymerase Reaction buffer dNTP mix MgCl2 Specific primer set Mineral oil Agarose Electrophoresis buffer (TAE or TBE) Gel-loading buffer Size marker Ethidium bromide Deionized water Terminator reaction mix Template-specific primer Ethanol EDTA

Supplies and Equipments Programmable thermal controller Electroporesis tank for horizontal slab gel Power supply UV transilluminator Microcentrifuges, room temp Vortex mixer 1.5 ml snap cap tubes Pipettes of 10–20, 100–200 and 1000 µl Aerosol resistant tips (filter tips) for each pipettes 0.2 ml or 0.5 ml PCR tubes Sequencer Spin columns for purifying PCR products Spin columns for purifying extension products Computer and the Internet

samples are also required to be examined for eggs or proglottids or copro-antigens or copro-DNA.41 Additional samples may be obtained through chemotherapy. The second is for mass screening in endemic areas for detection of carriers based on both questionnaire on the history of expulsion of proglottids and fecal and serological examinations. The third is for screening of personnel working in farming or meat production companies or butchers. Samples requested to be examined are as follows: (i) proglottids expelled from the carriers, (ii) feces and (iii) blood or serum samples from suspected persons or from epidemiological study samples. (i) Proglottids are kept in some sealed containers for rapid examination. If proglottids are not ready for

rapid examination, they better be kept either in PBS or in ethanol (> 70%) or in 5–10% formalin. For molecular examination, samples are recommended not to be kept in formalin. However, specimens kept in formalin for short time are still ready for molecular identification.7,42,43 Therefore, proglottids on the slide glass may also be applicable for detection of DNA.31 (ii) Feces are recommended to be kept frozen if possible, or to be divided into two aliquots. One is kept fresh for immediate copro-antigen test or frozen. The other is kept in ethanol for molecular test. Copro-antigen test is the first choice for screening or detection of taeniasis carriers.57,99,100 If it becomes positive, the rest of sample is for copro DNA test for identification of the species.41 (iii) Blood or serum is kept in refrigerator or frozen until detection of specific antibodies. There is a report stressing species specific serology to detect and differentiate taeniasis cases.101 However, it is still unclear without crucial evaluation with sera from carriers of T. saginata or T. asiatica. By contrast, there is unpublished serological tool to detect antibodies to genus specific antigens (Nakao et al. unpublished data). Serology for detection of specific antibodies in blood samples is not widely applied. It is, therefore, essential to confirm proglottids of Taenia species for detection of taeniasis cases.23

62.2.3  Detection Procedures 62.2.3.1  Multiplex PCR Compared to PCR-RFLP and BESS T-base analysis,102 multiplex PCR41 is a simpler and more rapid method. Multiplex PCR41 is useful for differentiation among not only the species of human Taenia but also two genotypes of T. solium. Specific primer set. Primers are derived from cox1, i.e., TsagMpF: TTGATTCCTTCGATGGCTTTTCTTTTG TasiMpF: ACGGTTGGATTAGATGTTAAGACTA TsolAmMpF: GGTAGATTTTTTAATGTTTTCTTTA

846

Molecular Detection of Foodborne Pathogens

TsolAsiMpF: TTGTTATAAATTTTTGATTACTAAC TaeniaMpR: GACATAACATAATGAAAATG

T. saginata

T. asiatica

T. solium Amer/Africa

T. solium Asian

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 M

Procedure:

1.5

(1) Prepare a master mix according as follows.

1.0 Component

Volume/Reaction

Final Concentration

10 × PCR buffer* dNTP mixture**

2.5 µl

1 × 

2.0 µl

200 µM of each dNTP

Primer TsagMpF (5 µM)

1.25 µl

0.25 µM

Primer TasiMpF (5 µM)

1.25 µl

0.25 µM

Primer TsolAmMpF (5 µM)

1.25 µl

0.25 µM

Primer TsolAsiMpF (5 µM)

1.25 µl

0.25 µM

Primer TaeniaMpR (5 µM) Taq polymerase

1.25 µl

0.25 µM 1 unit/reaction

Template DNA

 < 1 µg Variable

Deionized water Total volume

0.1 µl

Volume 25 µl

*Contains 15mM MgCl2, **2.5 mM of each.



Note: In general, it is assumed that the concentration and proportion of forward-to-reverse primers are important in the multiplex PCR. Although the optimal ratio of these primers was shown in the original paper41, the above master mix produces clear PCR products in our laboratory. Nonetheless, an ideal reaction composition should be determined in each laboratory. The master mix contains all the components needed for PCR except the template DNA. Prepare a volume of master mix 10% greater than that required for the total number of PCR assays to be performed. A negative control (without template DNA) and positive controls (T. saginata, T. asiatica and two genotypes of T. solium) should always be included. (2) Mix the master mix thoroughly and dispense appropriate volumes into PCR tubes. (3) Add template DNA (< 1 µg/reaction) to the individual tubes containing the master mix. (4) Proceed directly to next step, when using a thermal controller with a heated lid. Otherwise, overlay each reaction with mineral oil. (5) Program the thermal cycler. In the original paper by Yamasaki et al.,41 the multiplex PCR protocols consisted of 35 cycles of denaturation (30 sec at 94°C), annealing (30 sec at 60°C), and extension (90 sec at 72°C), plus one cycle of 5 min at 72°C. In our experiments, however, very weak or no band was obtained with this PCR protocol. On the other hand, nonspecific bands that gave unreliable results were observed at lower annealing temperature. In our case, when protocols consisted of predenaturation (5 min at 94ºC), 35 cycles of denaturation (30 sec at 94°C), annealing (30 sec at 57°C), and extension (70 sec at 72°C), plus one cycle of 7 min at 72°C, the most preferable result was obtained. PCR program should be adjusted to the thermal cycler used.

0.5

827 bp

269 bp

720 bp

984 bp

Figure 62.3  Multiplex PCR for differentiation of three human taeniid species including two genotypes of T. solium. (Modified from Yamasaki, H. et al., J. Clin. Microbiol., 42, 548, 2004.)

(6) Place the PCR tubes in the thermal cycler and start the cycling program. After amplification, samples can be stored overnight at 2–8°C or at –20°C for longer storage. (7) Load the PCR product onto an agarose gel (1%) with prior addition of gel-loading buffer. (8) Close the lid of the gel tank and attach the electrical leads so that the DNA will migrate toward the anode. Run the gel until the dye (bromophenol blue and/or xylene cyanol) has migrated the appropriate distance through the gel. (9) Turn off the electric current. If ethidium bromide is present in the gel and electrophoresis buffer, examine the gel by ultraviolet light and photograph. Otherwise, stain the gel with ethidium bromide (0.5 µg/ml) then take a photograph (Figure 62.3). 62.2.3.2 Sequencing of the PCR Product For epidemiological or retrospective studies, sequence data from the individual worm is very useful. Target gene. In the case of human taeniid cestodes, cox1 seems to be suitable. The length of cox1 in taeniid cestodes is relatively long (about 1620 bp), and the primer set which can amplify complete sequence of cox1 has been designed (Cox1/F, Cox1/R).39 When it is expected that worms collected from human may be taeniid species and that large molecule of DNA can be extracted from these worms, this primer set is suitable for the PCR amplification. Sequences of these primers are as follows: Cox1/F: GTTATGTTAGACTAGATGTTTTCA Cox1/R: TCCACTAAGCATAATGCAAAAGGC In the case of formalin-fixed samples or histopathological specimens, it may be difficult to amplify more than 1-kb DNA fragments, because of the fragmentation of DNA. Species of cysticerci recovered from human or domestic animals is not always human Taenia species. In these cases, Cox1/F/ Cox1/R may not function, and universal primer sets which amplify shorter fragments may be useful. The universal primer set, pr-a/pr-b, can amplify 444 bp fragment of cox1.61 In our experience, this primer set is applicable to all species

847

Taenia

of cestodes and trematodes examined, and also amplifies the cox1 fragment of some nematode species. Sequences of pr-a/ pr-b are as follows: pr-a: TGGTTTTTTGTGCATCCTGAGGTTTA pr-b: AGAAAGAACGTAATGAAAATGAGCAAC Procedure: (1) Prepare a master mix according as follows. Component

Volume/Reaction

Final Concentration

10 × PCR buffer* dNTP mixture**

2.5 µl

1 × 

2.0 µl

200 µM of each dNTP

Forward primer (5 µM)

1.25 µl

0.25 µM

Reverse primer (5 µM) Taq polymerase

1.25 µl

0.25 µM 1 unit/reaction

Template DNA

 < 1µg Variable

Deionized water Total volume

0.1 µl

25 µl

*Contains 15 mM MgCl2, **2.5 mM of each.

(2) Amplify the DNA fragment of the target gene. PCR protocols for Cox1/F/Cox1/R consisted of 30 cycles of denaturation (30 sec at 94°C), annealing (30 sec at 56°C), and extension (90 sec at 72°C), plus one cycle of 7 min at 72°C. PCR protocols for pr-a/pr-b consisted of predenaturation (5 min at 94 ºC), 30 cycles of denaturation (50 sec at 94°C), annealing (45 sec at 42°C), and extension (90 sec at 72°C), plus one cycle of 7 min at 72°C. (3) Purify the templates. For optimal results, purify the PCR product before sequencing. In general, any method that removes dNTPs and primers such as MinElute PCR Purificaticn Kit and Centricon-100 columns should work. (4) Perform cycle sequencing. Cycle sequencing provides the most reproducible results for sequencing PCR templates. Although PCR fragments can be difficult to denature with traditional sequencing methods, cycle sequencing provides several chances to denature and extend the template, ensuring adequate signal in the sequencing reaction. In the following, the example using BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems) is shown. The reaction mixtures: Reagent

Quantity

Terminator Ready Reaction Mix 8.0 µl Template 10–40 ng for Cox1/F/Cox1/R 3–10 ng for pr-a/pr-b Primer 3.2 pmol Deionized water Variable Total volume 20 µl

Place the tubes in a thermal cycler and set to the correct volume. Cycle sequencing protocols consisted

of predenaturation (5 min at 94ºC), 35 cycles of denaturation (10 sec at 96°C), annealing (5 sec at 50°C), and extension (4 min at 60°C). (5) Purify extension products: The best results are obtained when unincorporated dye terminators are completely removed prior to electrophoresis. Excess dye terminators in sequencing reactions obscure data in the early part of the sequence and can interfere with base-calling. Spin Column such as CentriSep can be used for purification. (6) Electrophorese. Purified extension products are electrophoresed by the sequencing instrument such as ABI PRISM 3100 Genetic Analyzer.

62.3 Conclusions and Future Perspectives Through local endemic areas in Asia and the Pacific, the risk factor for contamination of the environment is the butcher who sell meat and viscera in local markets.23,24 Even in developed countries, all personnel who are involved in production of meat including owners of the pastures, personnel working in the pasture are to be checked for such tapeworms. Production of safe meat under sustainable public health education with safe sanitation may be the only way to ensure the relative safety for the meat we consume.23,24 For improved evaluation of polymorphism of mitochondrial DNA of human taeniid cestodes, more detailed molecular probes to differentiate intraspecies variations might become important in order to do retrospective studies of the areas where domestic animals or humans had chances to be infected with these cestodes, either eggs or metacestodes. Then, if it is possible, the long DNA sequence from each specimen is to be analyzed to obtain additional information to track back where the patients were exposed to eggs or cysticerci using the retrospective specimens.48 For this purpose, cox1 seems to be suitable. The length of cox1 in taeniid cestodes is relatively long (about 1620 bp), and the primer set which can amplify complete sequence of cox1 has been designed.39 In the case of formalin-fixed samples or histopathological specimens, it may be difficult to amplify more than 1-kb DNA fragments, because of the fragmentation of DNA.7,50,51 In that case, we can select another primer set which amplify a shorter fragment such as P1/P2.103 Application of molecular approaches together with classic morphology is crucial for identification of the species or strains of parasites responsible for foodborne diseases.

Acknowledgments The projects summarized in this review had financial support from the Japan Society for the Promotion of Science (JSPS) (14256001, 17256002) and from the Ministry of Education, Japan (Kagaku-Gijutsu Shinko Choseihi 2003-2005) and JSPS-Asia/Africa Science Platform Fund (2006-2008) to A. Ito, from JSPS (18790289) to Y. Sako, and from JSPS (16500277, 18406008) to M. Okamoto.

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References











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Molecular Detection of Foodborne Pathogens 22. Bowles, J. and McManus, D.P. Genetic characterization of the Asian Taenia, a newly described taeniid cestodes of humans. Am. J. Trop. Med. Hyg., 50, 33, 1994. 23. Wandra, T. et al. High prevalence of Taenia saginata and status of Taenia solium cysticercosis in Bali, Indonesia, 2002–2004. Trans. R. Soc. Trop. Med. Hyg., 100, 346, 2006. 24. Wandra, T. et al. Taeniasis and cysticercosis in Bali and north Sumatra, Indonesia. Parasitol. Int., 55, S155, 2006. 25. Wandra, T. et al. Current situation of taeniasis and cysticercosis in Indonesia. Trop. Med. Health, 35, 323, 2007. 26. Schantz, P.M. et al. Neurocysticercosis in an orthodox Jewish community in New York City. N. Engl. J. Med., 327, 692, 1992. 27. Al Shahrani, D. et al. First case of neurocysticercosis in Saudi Arabia. J. Trop. Pediatr., 49, 58, 2003. 28. Hira, P.R. et al. Cysticercosis: imported and autochthonous infections in Kuwait. Trans. R. Soc. Trop. Med. Hyg., 98, 233, 2004. 29. Ito, A. et al. Towards the international collaboration for detection, surveillance and control of taeniasis/cysticercosis and echinococcosis in Asia and the Pacific. Southeast Asian J. Trop. Med. Public Health, 37 (Suppl 3), 82, 2006. 30. Li, T., et al. Taeniasis/cysticercosis in a Tibetan population in Sichuan province, China. Acta Trop., 100, 223, 2006. 31. Ananthapuruti, M.T. et al. Sympatric occurrence of Taenia solium, T. saginata, and T. asiatica, Thailand. Emerg. Infect. Dis., 13, 1413, 2007. 32. Lloyd, S. Cysticercosis and taeniosis Taenia saginata, Taenia solium, and Asian Taenia. In Zoonoses: Biology, Clinical Practice, and Public Health Control (eds by Palmer, S.R., Soulsby, L. and Simpson, D.I.H.), 635. Oxford University Press, New York, 1998. 33. Ito A. Welcome remarks and introduction to symposium on cestodes zoonoses in Asia and the Pacific. Southeast Asian J. Trop. Med. Public Health, 38 (Suppl 1), 115, 2007. 34. Flisser, A., Craig, P.S., and Ito, A. Cysticercosis and taeniosis Taenia saginata, Taenia solium, and Asian Taenia. In Zoonoses: Biology, Clinical Practice, and Public Health Control, 2nd edition (eds by Soulsby, E.J.L., Thorgerson, P). Oxford University Press, New York, in press. 35. Zarlenga, D.S., et al. Characterization and detection of a newly described Asian taeniid using cloned ribosomal DNA fragments and sequence amplification by the polymerase chain reaction. Exp. Parasitol., 72, 174, 1991. 36. Fan, In Studies on Taceniasis (1981–2002) (ed by Fan, P.C.). National Yangming University, Taipei, 109, 1992. 37. Simanjuntak G, M., et al. Taeniasis/cysticercosis in Indonesia as an emerging disease. Parasitol. Today, 13, 321, 1997. 38. Okamoto, M. et al. Molecular variation of Taenia solium in the world. Southeast Asian J. Trop. Med. Public Health, 32 (Suppl 2), 90, 2001. 39. Nakao, M. et al. A phylogenetic hypothesis for the distribution of two genotypes of the pig tapeworm Taenia solium worldwide. Parasitology, 124, 657, 2002. 40. Ito, A. et al. Mitochondrial DNA of Taenia solium: from basic to applied science. In Taenia solium Cysticercosis (eds by Singh, G., Prabhakar, S.), 47. CAB International, Oxon, 2002. 41. Yamasaki, H. et al. DNA differential diagnosis of taeniasis and cysticercosis by multiplex PCR. J. Clin. Microbiol., 42, 548, 2004. 42. Yamasaki, H. et al. Mitochondrial DNA diagnosis for taeniasis and cysticercosis. Parasitol. Int., 55, S81, 2006. 43. Yamasaki, H. et al. Significance of molecular-pathological diagnosis in cestodes zoonoses. Trop. Med. Health, 35, 307, 2007.

Taenia 44. Wandra, T., et al. Resurgence of cases of epileptic seizures and burns associated with cysticercosis in Assologaima, Jayawijaya, Irian Jaya, Indonesia, 1991–95. Trans. R. Soc. Trop. Med. Hyg., 94, 46, 2000. 45. Ito, A. et al. Cysticercosis/taeniasis in Asia and the Pacific. Vector-Borne Zoonotic Dis., 4, 95, 2004. 46. Ito, A., et al. Multiple genotypes of Taenia solium—ramifications for diagnosis, treatment and control. Acta Trop., 87, 95, 2003. 47. Campbell, G. et al. Genetic variation in Taenia solium. Parasitol. Int., 55, S121, 2006. 48. Sudewi, A.A.R. et al. Taenia solium cysticercosis in Bali, Indonesia: serology and mtDNA analysis. Trans. R. Soc. Trop. Med. Hyg., 102, 96, 2008. 49. Yamasaki, H., et al. Molecular identification of Taenia solium cysticercus genotype in the histopathological specimens. Southeast Asian J. Trop. Med. Public Health, 36 (Suppl 4), 131, 2005. 50. Yamasaki, H., et al. A case of intramuscular cysticercosis diagnosed definitively by mitochondrial DNA analysis of extremely calcified cysts. Parasitol. Int., 55, 127, 2006. 51. Yamasaki, H., et al. Mitochondrial DNA diagnosis for cestode zoonoses: application to formalin-fixed paraffin-­embedded ­tissue specimens. Southeast Asian J. Trop. Med. Public Health, 38 (Suppl 1), 166, 2007. 52. Almeida, C.R., et al. Taenia solium DNA is present in the cerebrospinal fluid of neurocysticercosis patients and can be used for diagnosis. Eur. Arch. Psychiatry Clin. Neurosci., 256, 307, 2006. 53. Ito, A. et al. Neurocysticercosis case with a single cyst in the brain showing dramatic drop in specific antibody titers within 1 year after curative surgical resection. Parasitol. Int., 48, 95, 1999. 54. Ohsaki, Y., et al. Neurocysticercosis without detectable ­specific antibody. Intern. Med., 38, 67, 1999. 55. Chung, J.Y., et al. Molecular determination of the origin of acephalic cysticercus. Parasitology, 130, 239, 2005. 56. Eom, K.S., et al. Identification of Taenia asiatica in China: molecular, morphological, and epidemiological analysis of a Luzhai isolate. J. Parasitol., 88, 758, 2002. 57. Allan, J.C. and Craig, P.S. Coproantigens in taeniasis and echinococcosis. Parasitol. Int., 55, S75, 2006. 58. Nunes, C.M., et al. Taenia saginata: differential diagnosis of human taeniasis by polymerase chain reaction-restriction fragment length polymorphism assay. Exp. Parasitol., 110, 412, 2005. 59. Notomi, T. et al. Loop-mediated isothermal amplification of DNA. Nucleic Acids Res., 28, E63, 2000. 60. Nkouawa, A. et al. Loop-mediated isothermal amplification method for differentiation and rapid detection of Taenia species. J. Clin. Microbiol., 47, 168, 2009. 61. Okamoto, M. et al. Phylogenetic relationships within Taenia taeniaeformis variants and other taeniid cestodes inferred from the nucleotide sequence of the cytochrome c oxidase subunit I gene. Parasitol. Res., 81, 451, 1995. 62. Theis, J.H. et al. DNA-confirmed Taenia solium cysticercosis in black bears (Urus americanus) from California. Am. J. Trop. Med. Hyg., 55, 456, 1996. 63. Takayanagui, O.M. and Odashima, N.S. Clinical aspects of neurocysticercosis. Parasitol. Int., 55, S111, 2006. 64. Tsang, V.C., Brand, J.A. and Boyer, A.E. An enzyme-linked immunoelectro-transfer blot assay and glycoprotein antigens for diagnosing human cysticercosis. J. Infect. Dis., 159, 50, 1989.

849 65. Wilson, M. et al. Clinical evaluation of the cysticercosis enzyme-linked immunoelectrotransfer blot in patients with neurocysticercosis. J. Infect. Dis., 164, 1007, 1991. 66. Ito, A. et al. Novel antigens for neurocysticercosis: simple method for preparation and evaluation for serodiagnosis. Am. J. Trop. Med. Hyg., 59, 291, 1998. 67. Chung, J.Y. et al. A recombinant 10-kDa protein of Taenia solium metacestodes specific to active neurocysticercosis. J. Infect. Dis., 180, 1307, 1999. 68. Sako, Y. et al. Molecular characterization and diagnostic value of Taenia solium low-molecular-weight antigen gene. J. Clin. Microbiol., 38, 4439, 2000. 69. Green, R.M. et al. Taenia solium: molecular cloning and serological evaluation of 14- and 18-kDa related, diagnostic antigens. J. Parasitol., 86, 1001, 2000. 70. Hancock, K. et al. Characterization of the 8-kilodalton antigens of Taenia solium metacestodes and evaluation of their use in an enzyme-linked immunosorbent assay for serodiagnosis. J. Clin. Microbiol., 41, 2577, 2003. 71. Hancock, K. et al. Characterization and cloning of T24, a Taenia solium antigen diagnostic for cysticercosis. Mol. Biochem. Parasitol., 147, 109, 2006. 72. Sato, M.O. et al. Evaluation of tongue inspection and serology for diagnosis of Taenia solium cysticercosis in swine: usefulness of ELISA using purified glycoproteins and recombinant antigen. Vet. Parasitol., 111, 309, 2003. 73. Sato, M.O. et al. Evaluation of purified Taenia solium glycoproteins and recombinant antigens in the serologic detection of human and swine cysticercosis. J. Infect. Dis., 194, 1783, 2006. 74. Ito, A. et al. ELISA and immunoblot using purified glycoproteins for serodiagnosis of cysticercosis in pigs naturally infected with Taenia solium. J. Helminthol., 73, 363, 1999. 75. Wandra, T. et al. Taenia solium cysticercosis, Irian Jaya, Indonesia. Emerg. Infect. Dis., 9, 884, 2003. 76. Furrows, S.J. et al. Lack of specificity of a single positive 50-kDa band in the electroimmunotransfer blot (EITB) assay for cysticercosis. Clin. Microbiol. Infect., 12, 459, 2006. 77. Handali, S., et al. Porcine antibody responses to Taenia solium antigens RGP50 and STS18VAR1. Am. J. Trop. Med. Hyg., 71, 322, 2004. 78. Scheel. C.M. et al. Serodiagnosis of neurocysticercosis using synthetic 8-kDa proteins: comparison of assay formats. Am. J. Trop. Med. Hyg., 73, 771, 2005. 79. Arrunda, G.C., et al. Evaluation of Taenia solium and Taenia crassiceps cysticercal antigens for the serodiagnosis of neurocysticercosis. Trop. Med. Internat. Health, 10, 1005. 2005. 80. Espindola, N.M. et al. Cysticercosis immunodiagnosis using 18- and 14-kilodalton proteins from Taenia crassiceps cysticercus antigens obtained by immunoaffinity chromatography. J. Clin. Microbiol., 43, 3178, 2005. 81. Oliveira, H.B. et al. Application of Taenia saginata metacestodes as an alternative antigen for the serological diagnosis of human neurocysticercosis. Parasitol. Res., 101, 1007, 2007. 82. Sciutto, E. et al. Improvement of the synthetic tri-peptide vaccine (S3Pvac) against porcine Taenia solium cysticercosis in search of a more effective, inexpensive and manageable vaccine. Vaccine, 25, 1368, 2007. 83. Ito, A. Serologic and molecular diagnosis of zoonotic larval cestode infections. Parasitol. Int., 51, 221, 2002. 84. Ito, A. and Craig, P.S. Immunodiagnostic and molecular approaches for the detection of taeniid cestode infections. Trends Parasitol., 19, 377, 2003.

850 85. Ito, A. and Craig, P.S. Response to Dorny et al.: immunodiagnostic approaches for detecting Taenia solium. Trends Parasitol., 20, 260, 2004. 86. Harrison, L.J. et al. Specific detection of circulating surface/ secreted glycoproteins of viable cysticerci in Taenia saginata cysticercosis. Parasite Immunol., 11, 351, 1989. 87. Brandt, J. et al. A monoclonal antibody based ELISA for the detection of circulating excretory-secretory antigens in Taenia saginata cysticercosis. Int. J. Parasitol., 22, 471, 1993. 88. Dorny, P. et al. A Bayesian approach for estimating values for prevalence and diagnostic test characteristics of porcine cysticercosis. Int. J. Parasitol., 569, 2004. 89. Del Bruto, O.H., et al. Proposed diagnostic criteria for neurocysticercosis. Neurology, 57, 177, 2001. 90. Schantz, P.M. Progress in diagnosis, treatment and elimination of echinococcosis and cysticercosis. Parasitol. Int., 55, S7, 2006. 91. Fan, P.C. et al. Eating habits of East Asian people and transmission of taeniasis. Acta Trop., 50, 305, 1992. 92. Fan, P.C. et al. Experimental infection of Taenia saginata asiatica (Guizhou strain) in domestic animals and morphological characteristics. Chinese J. Parasitol., 14, 45, 2001. 93. Nakaya, K. et al. Usefulness of severe combined immunodeficiency (scid) and inbred mice for studies of cysticercosis and echinococcosis. Parasitol. Int., 55, S91, 2006. 94. Ito, A. et al. Human Taenia eggs develop into cysticerci in scid mice. Parasitology, 114, 85, 1997. 95. Ito, A. and Ito, M. Human Taenia in severe combined immunodeficiency (scid) mice. Parasitol. Today, 15, 64, 1999. 96. Nakao, M., et al. Isolation of polymorphic microsatellite loci from the tapeworm Echinococcus multilocularis. Infect, Genet, Evol., 3, 159, 2003. 97. Bart, J.M., et al. EmsB, a tandem repeated multi-loci microsatellite, new tool to investigate the genetic diversity of Echinococcus multilocularis. Infect, Genet, Evol., 6, 390, 2006. 98. Knapp, J, et al. Assessment of use of microsatellite polymorphism analysis for improving spatial distribution tracking of Echinococcus multilocularis. J. Clin. Microbiol., 45, 2943, 2007. 99. Allan, J.C. et al. Coproantigen detection for the immunodiagnosis of echinococcosis and taeniasis in dogs and humans. Parasitology, 104, 347, 1992.

Molecular Detection of Foodborne Pathogens 100. Allan, J.C. et al. Field trial of diagnosis of Taenia solium taeniasis by coproantigen enzyme-linked immunosorbent assay. Am. J. Trop. Med. Hyg., 54, 352, 1996. 101. Wilkins, P.P. et al. Development of a serologic assay to detect Taenia solium taeniasis. Am. J. Trop. Med. Hyg., 60, 199, 1999. 102. Yamasaki, H. et al. DNA differential diagnosis of human taeniid cestodes by base excision sequence scanning ­thymine-base reader analysis with mitochondrial genes. J. Clin. Microbiol., 40, 3818, 2002. 103. Rishi, A.K. and McManus, D.P. DNA probes which unambiguously distinguish Taenia solium from T. saginata. Lancet, 2 (8570), 1275, 1987. 104. Flisser, A. et al. Specific detection of Taenia saginata eggs by DNA hybridization. Lancet, 2 (8625), 1429, 1988. 105. Harrison, L.J., Delgado, J. and Parkhouse, R.M. Differential diagnosis of Taenia saginata and Taenia solium with DNA probes. Parasitology, 100, 459, 1990. 106. Chapman, A. et al. Isolation and characterization of speciesspecific DNA probes from Taenia solium and Taenia saginata and their use in an egg detection assay. J. Clin. Microbiol., 33, 1283, 1995. 107. González, L.M. et al. PCR tools for the differential diagnosis of Taenia saginata and Taenia solium taeniasis/­cysticercosis from different geographical locations. Diagn. Microbiol. Infect. Dis., 42, 243, 2002. 108. Gasser, R.B., Zhu, X. and Woods, W. Genotyping Taenia tapeworms by single-strand conformation polymorphism of mitochondrial DNA. Electrophoresis, 20, 2834, 1999. 109. Vega, R. et al. Population genetic structure of Taenia solium from Madagascar and Mexico: implications for clinical profile diversity and immunological technology. Int. J. Parasitol., 33, 1479, 2003. 110. Dias, A.K. et al. Taenia solium and Taenia saginata: identification of sequence characterized amplified region (SCAR) markers. Exp. Parasitol., 117, 9. 2007. 111. González, L.M. et al. Differential diagnosis of Taenia saginata and Taenia solium infection by PCR. J. Clin. Microbiol., 38, 737, 2000. 112. von Nickisch-Rosenegk, M., Silva-Gonzalez, R. and Lucius, R. Modification of universal 12S rDNA primers for specific amplification of contaminated Taenia spp. (Cestoda) gDNA enabling phylogenetic studies. Parasitol. Res., 85, 819, 1999. 113. Nakao, M. et al. Mitochondrial genetic code in cestodes. Mol. Biochem. Parasitol., 111, 415, 2000.

63 Trichinella

Edoardo Pozio and Giuseppe La Rosa National Institute of Health

Contents 63.1. Introduction..................................................................................................................................................................... 851 63.1.1. Taxonomy.......................................................................................................................................................... 851 63.1.1.1. The Encapsulated Clade................................................................................................................... 851 63.1.1.2. The Nonencapsulated Clade............................................................................................................. 853 63.1.2. Biology.............................................................................................................................................................. 853 63.1.3. Pathogenesis...................................................................................................................................................... 853 63.1.4. Medical Importance.......................................................................................................................................... 854 63.1.5. Methods for Diagnosing Trichinella Infection.................................................................................................. 854 63.2. Methods........................................................................................................................................................................... 856 63.2.1. Sample Collection and Preparation................................................................................................................... 856 63.2.1.1. Reference Material............................................................................................................................ 856 63.2.1.2. Hosts and Preferential Muscles......................................................................................................... 856 63.2.1.3. Parasite Isolation and Preservation................................................................................................... 856 63.2.1.4. DNA Purification from Single Larvae.............................................................................................. 857 63.2.2. Detection Procedures........................................................................................................................................ 858 63.2.2.1. Multiplex PCR.................................................................................................................................. 858 63.2.2.2. PCR-RFLP to Distinguish between Trichinella britovi and Trichinella T9.................................... 858 63.2.2.3. PCR to Distinguish between Trichinella britovi and Trichinella T8............................................... 859 63.2.2.4. SSCP................................................................................................................................................. 860 63.3. Conclusions and Future Perspectives.............................................................................................................................. 860 References.................................................................................................................................................................................. 861

63.1  Introduction Nematode worms belonging to the genus Trichinella are the etiological agent of the zoonosis trichinellosis, which is the appropriate term for the human disease also known as “trichinosis” or “trichiniasis.” On all continents but Antarctica, these parasites are widespread among wildlife, and in many countries they are widespread among domestic pigs.1 Human infection occurs following the consumption of raw or undercooked meat and meat products from different animals (e.g., pork, horse, game). The main distinguishing feature of the life cycle of these parasites is that two generations occur in the same host. The first generation (from the infective larva to the adult) is present in the gut, and the second generation (from the newborn larva to the infective larva) is present in the cell of striated muscles, which is modified by the larva (referred to as a “nurse cell”).2

63.1.1 Taxonomy Until 1972, Trichinella spiralis was the only known species in the Trichinella genus, and for this reason it was the only species considered in most of the studies conducted up through the 1990s. At present, there are eight known species

and four additional genotypes that have not been characterized at the species level. Morphologically, the individual species and genotypes cannot be distinguished, yet they can be categorized into two clades based on whether or not the nurse cells in the host’s muscles are surrounded by a collagen capsule, which is important for the long-term survival of the larva in decaying muscle tissue (i.e., when the larva is not protected by the host). If the collagen capsule is present, then the species/ genotype belongs to the encapsulated clade, which consists of five species and the four genotypes; otherwise it belongs to the non-encapsulated clade. However, the non-encapsulated larvae induce the same modifications as the encapsulated larvae to the muscle cell, though the amount of collagen produced is very low. A brief description of the encapsulated and non-encapsulated species and genotypes is provided below. 63.1.1.1 The Encapsulated Clade T. spiralis is the species that has been characterized the most, not only because it was the first to have been discovered but also because of its importance both as a cause of human disease and as a model for basic biological research, particularly in the field of immunology, due to the relatively high prevalence of infection in domestic and wild animals and the 851

852

Molecular Detection of Foodborne Pathogens

marked infectivity for laboratory animals. The species originated in East Asia,3 and its worldwide dissemination was facilitated by the European colonization of the Americas, New Zealand, Hawaii, and Egypt from the 16th century to the 20th century (Table 63.1).4 The predominant hosts are domestic and wild swine (Sus scrofa), synanthropic animals, other domestic animals, and diverse species of wild carnivores. T. spiralis is the etiological agent of most human infections worldwide. It is more pathogenic than the other species and genotypes because the female produces a higher number of newborn larvae5 and because it induces a greater immune response in humans.6,7 Trichinella nativa is the etiological agent of infection in ­sylvatic carnivores in frigid zones of Asia (China, Kazakhstan, Mongolia, Russia), North America (Canada, Greenland, and the USA) and Europe (Estonia, Finland, Lithuania, Norway, Russia, and Sweden) (Table 63.1). The southern boundary of distribution has been tentatively identified to lie between the isotherms –5° and –4°C in January. The main hosts are polar, brown, and black bears; wolverines; raccoons; lynxes; wolves; foxes; and walruses.8,9 One of this species’ biological characteristic is the larva’s ability to survive in frozen muscles of carnivores for up to 5 years.9 Human infection has been reported in frigid zones of Canada, Greenland, Siberia, and Kamchatka10–12 and among hunters from Europe and the

USA who consume raw or undercooked meat from bears hunted in arctic or subarctic regions.13 The genotype Trichinella T6 is closely related to T. nativa and has been detected in several regions of Canada (British Columbia, Ontario, Manitoba, and Nunavut) and the USA (Alaska, Montana, Idaho, and Pennsylvania).14 The larvae of this genotype, like those of T. nativa, can survive in frozen muscles of carnivores for long periods of time.9 Trichinella britovi is the etiological agent of infection in sylvatic carnivores (e.g., foxes, wolves, badgers, stone martens, jackals, bears, lynxes, cats, and dogs) and omnivores (in most cases the wild boar) in temperate areas of Europe and Asia (from the Iberian Peninsula to Kazakhstan) and in North and West Africa (Table 63.1). The northern boundary of distribution appears to be between the isotherms –6° and –5° C in January. The larvae of T. britovi can survive in frozen muscles of carnivores for up to 11 months and in frozen muscles of swine for up to 3 weeks, which, however, is a shorter period of time compared to T. nativa larvae.9 Human infection caused by the consumption of free-ranging pigs, game, and horse meat has been documented in Europe, Asia, and Africa north of the Sahara.15–19 The clinical course is relatively benign, and no deaths have been documented.20 The genotype Trichinella T8 is related to T. britovi. It has been detected in sylvatic carnivores of South Africa and

Table 63.1 Principal Features of Trichinella Species and Genotypes and Amplicon Sizes by PCR-RFLP Trichinella Species or Genotype

Distribution

Trichinella T8 T. murrelli Trichinella T9 T. nelsoni Trichinella T12

Cosmopolitana Arctic and subarctic areas of Holoarctic regionb Canada and United States Temperate areas of Palearctic regionc, northern and western Africa South Africa and Namibia Temperate areas of Nearctic region Japan Ethiopic region Argentina

T. pseudospiralis

Cosmopolitan

T. papuae

Papua New Guinea Thailand Ethiopia, Mozambique, South Africa, Zimbabwe

T. spiralis T. nativa Trichinella T6 T. britovi

T. zimbabwensis

Cycle Encapsulated Domestic and sylvatic Sylvatic

b

Amplicon Size by PCR-RFLP (Mse I)

22, 70, 126, 201 bp 22, 70, 327 bp

Sylvatic Sylvatic, seldom domestic

Swine, rats, seldom carnivores Terrestrial and marine carnivores Carnivores Carnivores, seldom swine

Sylvatic Sylvatic Sylvatic Sylvatic Sylvatic

Carnivores Carnivores Carnivores Carnivores, seldom swine Carnivores

22, 62, 64, 70, 201 bp 92, 126, 201 bp 92, 327 bp 62, 64, 70, 223 bp Unknown

Mammals and birds

419 bpd Unknowne Unknown

Non-encapsulated Sylvatic, seldom domestic

This species has not been detected in arctic regions. The isotherm –4°C in January is the southern limit of distribution. c The isotherm –6°C in January is the northern limit of distribution. d Palearctic region. e Nearctic and Australian (Tasmania) regions. Source: Nagano, I. et al., Int. J. Parasitol., 29, 1113, 1999. a

Hosts

Sylvatic, seldom domestic Sylvatic and domestic

Swine, Saltwater crocodiles Nile crocodiles, Nile monitor lizards, lion

22, 70, 327 bp 22, 62, 64, 70, 201 bp

Unknown

853

Trichinella

Namibia (Table 63.1).21 No human infections have been documented to date. Trichinella murrelli is the etiological agent of infection in wild carnivores in temperate areas of North America. The isotherm –6°C in January seems to be the northern boundary of distribution (Table 63.1). Because this species shows very low infectivity to swine and rats, it does not pose a risk for domestic pigs.22 Human infection has occurred for the consumption of bear meat in the USA and of horse meat exported from the USA to France.1 The genotype Trichinella T9, which is phylogenetically related to T. murrelli, has been detected in wild carnivores in Japan.23 No human infections have been documented. Trichinella nelsoni is the etiological agent of infection of wild carnivores (e.g., spotted and striped hyenas, lions, leopards, servals, bat eared foxes, and cheetahs) in East Africa, from Kenya to South Africa (Table 63.1).24 This species occasionally infects wild swine (e.g., warthogs and bush pigs), whose meat can be the source of human infection. Infection has never been detected in domestic pigs. To date, fewer than 100 human infections have been documented. This species shows low pathogenicity in humans, and deaths have only been reported in persons with more than 4,000 larvae per gram of muscle.25,26 Trichinella T12 is a newly discovered genotype, which was detected in a mountain lion (Puma concolor) from Trapalco (66°59′ W, 39°34′ S) (Patagonia, Río Negro, Argentina) (Table 63.1). This taxon is considered as a separate genotype based on the molecular structure of two noncoding sequences (expansion segment V, 5S ribosomal DNA intergenic spacer region) and one coding sequence (cytochrome c-oxidase subunit I; CO I), which are different from those of the other 11 species/genotypes.27 63.1.1.2 The Non-encapsulated Clade Trichinella pseudospiralis is a cosmopolitan species which infects both domestic and wild mammals; it is the only species that infects birds, both wild carnivorous and omnivorous (Table 63.1). Three populations, which can be distinguished on a molecular basis, have been detected: one population in Europe and Asia, one in North America, and one in Tasmania.28 T. pseudospiralis has been found in 14 mammal species and seven bird species;29 however, relatively few bird species have been examined, and the number of examined birds has also been low. In humans, the clinical severity of infection can range from moderate to severe; one death has been reported.30–33 Trichinella papuae has been detected in Papua New Guinea (PNG) and Thailand (Table 63.1).34–36 It can infect both mammals (i.e., wild pigs) and reptiles (i.e., saltwater crocodiles and monitor lizards). Wild pigs have been implicated in human infection.37 In PNG, it was reported that a high percentage of persons who consume wild pig were serologically positive for Trichinella antigens yet had not clinical signs of the disease, suggesting that they have frequent contact with this parasite.37 Trichinella zimbabwensis has been detected in farmed Nilecrocodiles of Zimbabwe38 and Ethiopia, in wild

crocodiles of Mozambique and South Africa, and in wild monitor ­lizards of Zimbabwe (Table 63.1).39 In laboratory conditions, as well as in nature, this species can infect mammals.

63.1.2  Biology A peculiarity of the biology of Trichinella nematodes is that two generations (i.e., adult worms and newborn larvae) are present in the host. Female worms embedded in the intestinal mucosa give birth to larvae, which migrate directly into the lymphatic vessels; they then enter the blood vessels and are spread throughout the entire body. Once the larvae reach the striated muscle cells, they penetrate the cell using a stiletto apparatus and lytic enzymes; this occurs in all striated muscles except for those of the heart. After about 15 days, the larvae develop to the L1 infective stage (at which point they are referred to as “muscle larvae”). In the muscle cells, larvae can survive for years (e.g., for over 20 years in polar bears and for up to 40 years in humans). When a new host ingests infected muscle tissue, the cells are digested in the stomach, releasing the larvae; when the larvae reach the duodenum, they penetrate the villa and within 2 days undergo four moults, developing to the adult stage. Males and females copulate, and from 6–7 days post infection (d.p.i.), the females begin to produce newborn larvae (i.e., the new generation), which continue to be produced for at least 1 or 2 weeks, or more, depending on the host’s immune response at the gut level, which results in the expulsion of adult worms. All Trichinella species and genotypes have a sylvatic cycle (i.e., natural transmission occurs among wildlife), though T.  spiralis is primarily transmitted by a domestic cycle involving domestic pigs, synanthropic rats and, on rare occasions, horses.1 A domestic cycle involving pigs and rats has also been reported for T. britovi and T. pseudospiralis, though rarely.1 The prevalence of infection among carnivorous mammals (e.g., foxes, wolves, racoon dogs, lynxes, bears, and mustelids) ranges from 0.01% to 90%, depending on the animal’s habitat and the extent of human influence.40,41 Given that Trichinella infection in sylvatic and domestic animals is mainly asymptomatic, even when thousands of larvae per gram of muscle tissue are present, there are no clinical signs that can help to identify infected animals.

63.1.3  Pathogenesis Humans acquire trichinellosis by consuming raw or undercooked meat from pigs, horses, dogs, and game animals (e.g., wild boar, fox, jackal, badger, bear, and walrus); the presence of Trichinella in domestic or wild animals is not in itself sufficient for transmission to humans. The clinical picture and prognosis of human infection depend on the number of infective larvae ingested, the Trichinella species, and the person’s immune response.42 About 0.2% of cases result in death, though the percentage has been reported to range from 0.1% to 1.0%. Death is usually due to respiratory or cardiovascular complications or neurological disorders.43

854

The pathological picture of infection is complex because, as mentioned, there exist two generations in the host: adults, which induce pathological alterations in the intestine; and larvae, which, during their development, induce alterations in the muscle. The extent of the immunological response is proportional to the number of larvae ingested, and it greatly depends on individual host type I hypersensitivity, which plays a major role in the early intestinal stage of the disease. The adult worms induce a severe intestinal pathology, including villus atrophy and crypt and goblet cell hyperplasia. This intestinal pathology is directly correlated with the inflammatory response inducing the spontaneous expulsion of adult worms (referred to as “self-expulsion”). In this process, the antibody production and cytokines produced by Th-2 cells play a major role. The activation of the Th-2 cytokines induces the stem cell factor (SCF) production, followed by the SCF differentiation, preferentially along the goblet cell lineage.44 Unlike other muscle parasites, Trichinella larvae transform the muscle cell, altering its metabolism and transforming the cell into a nurse cell, which allows the larva to survive and mature. The parasite elicits angiogenesis directly or indirectly by inducing changes in the nurse cells, which, in turn, stimulate the expansion of the network of blood vessels.44 This expansion could be explained by increased levels of vascular endothelial growth factor.45 In the heart muscle, the larvae and high levels of eosinophiles induce transient myocarditis, whereas damage to the central nervous system (CNS) may be far more severe. The pathogenesis of CNS lesions could be linked to the toxicity of eosinophiles for endothelial cells, leading to vasculitis and the infiltration of the pia-arachnoid with lymphocytes, macrophages, fibroblasts, and gitter cells. Nodules of glial cells and small hemorrhages may be present in the periventricular area and other areas of the white matter.

Molecular Detection of Foodborne Pathogens

petechiae, intraconjunctival hemorrhage, and hemorrhage of nail beds occur. These symptoms are accompanied by eosinophilia and usually by leukocytosis. This symptomatology is followed by pain in various muscle groups, which may restrict motility. Of these signs and symptoms, those considered to be the principal manifestations are: fever, eyelid, and periocular edema, and muscle pain.43 In severe cases, fever may be continuous and persist for up to 3 weeks; in mild cases, very mild fever is common. The intensity of periocular, eyelid, and facial edema depends on the host’s allergic reaction. The intensity of muscle pain reflects the severity of the disease. Pain occurs upon movement; spontaneous pain is less common, and according to some authors it occurs more frequently in persons with severe disease or as a result of complications in the form of thrombophlebitis. The symptoms gradually disappear during convalescence, and complete regression may be achieved with physical therapy.42 It has not been firmly established whether a chronic form of trichinellosis actually exists. Nonetheless, there have been reports of persons who, months or even years after the acute stage, continue to suffer from chronic pain, general discomfort, tingling, numbness, and excessive sweating, and who show signs of paranoia and a syndrome of persecution. The persistence of these symptoms has been more frequently observed among persons who had suffered from severe trichinellosis. There have been reports of impaired muscle strength, conjunctivitis, impaired co-ordination and the presence of IgG antibodies up to 10 years after infection, and live larvae have been detected in muscles up to 39 years after infection, yet without clinical signs or symptoms. All cases in which trichinellosis has been defined as “chronic” have been reported among persons who had not been treated in a timely manner.43

63.1.4 Medical Importance

63.1.5 Methods for Diagnosing Trichinella Infection

Worldwide, an average of 10,000 cases of clinical disease in humans has been estimated to occur each year, with a mortality rate of about 0.2%.2 However, in many countries the number of infections is underreported because appropriate serological tests are not available and because physicians are often not familiar with the disease. The acute stage of the disease corresponds with the phase in which the newborn larvae migrate from the lymphatic vessels and invade the muscle cells. In varying percentages of infected individuals, the acute stage is preceded by loose stools or diarrhea. These symptoms may be accompanied by flatulence, moderately intense abdominal pain, loss of appetite and, at times, vomiting.43 The signs and symptoms of the acute stage of trichinellosis develop and disappear at varying intervals and are of varying intensity. In most persons, the onset of the acute stage is sudden, with general weakness, chills, headache, fever (up to 40°C), excessive sweating and tachycardia. In the majority of cases, symmetrical eyelid and periocular edemas occur, and edema frequently affects the entire face.43 The blood vessels of the conjunctivae become inflamed and in some persons

In humans, which are the only host in which Trichinella can induce clinical disease, diagnosis is based on the following: (i) epidemiological data (i.e., the source of infection, the amount of infected meat ingested, the number of larvae present in the infected meat, and the number of cases in the epidemic focus); (ii) clinical evaluation (i.e., signs and symptoms and their clinical severity, which significantly affects the choice of treatment); and (iii) laboratory tests (i.e., immunodiagnosis and/or detection of larvae in a muscle biopsy using HCl-pepsin digestion). In animals, which show no clinical signs, the diagnosis is based exclusively on the detection of Trichinella larvae in muscle tissues using HCl-pepsin digestion. Given that morphological characters cannot be used to distinguish the taxon and that the biological and biochemical characters (e.g., allozymes) do not allow the species to be easily or rapidly identified, methods based on polymerase chain reaction (PCR) must be applied to the species or genotype determination. Identifying the species is extremely important in tracing the source of infection, in determining and predicting the clinical course of infection, in interpreting the clinical and laboratory data, in estimating the potential risk

Trichinella

for pigs, in establishing appropriate strategies for control and eradication, and in better understanding the epidemiology of the infection. PCR has proven to be important in simplifying the identification of Trichinella isolates from different host species and geographical regions (Table 63.1).46–61 The high sensitivity of PCR allows the species to be identified using a single larva, which is important because often only one larva is detected in samples from humans and animals. That the species can be identified with a single larva is also important because it allows different Trichinella species to be identified in the same host, a finding that has contributed to understanding the potential for gene flow between sympatric species or genotypes.1 PCR also allows the species to be identified based on a fraction of the larva (as small as 1/15th), which is important because in histological sections only a fraction of a larva is present. Molecular probes. The first molecular approaches for distinguishing among Trichinella species and genotypes were based on restriction enzyme analyses of genomic DNA and DNA probes. However, this method, in addition to being time consuming, requires a large number of parasites. This entails performing isolation and propagation in laboratory animals, which, however, can result in the loss of genetic variability and failure to identify mixed infections. Random amplified polymorphic DNA (RAPD). A plethora of PCR-derived methods have been developed to identify Trichinella larvae at the species or genotype level. These methods can be divided into those using arbitrary primers (RAPD) and those using DNA sequence-derived genotype-specific primers. The fingerprint pattern obtained with RAPD is a simple and effective tool for rapidly generating and detecting genetic markers; however, this method can be imprecise, producing bands that are not always reproducible and thus creating confusion in interpreting the results.49 Furthermore, problems associated with PCR signal reproducibility due to variations in DNA quality and contamination, reaction conditions and instrumentation, can generate substantial sample-to sample deviation.62 This, in turn, requires extensive controls to be performed with each analysis. A RAPD-derived approach has been developed to distinguish between T. spiralis and T. pseudospiralis.53 In this work, RAPD has been produced from the two species using purified genomic DNA and an arbitrary 10-mer primer. Two species-specific fragments have been isolated, sequenced, and used to design two primer sets that identify the two species using a standard PCR. In an expanded yet similar set of experiments, primer pairs with a high degree of specificity for T. spiralis, T. nativa, T. britovi, T. nelsoni, and T. pseudospiralis, have been generated.54 Problems and inconsistencies in using RAPD and associated technologies can led to a more deliberate approach towards designing genotype-specific PCR primers. Single-strand conformational polymorphism (SSCP). SSCP can be used to characterize and differentiate eight distinct taxa of Trichinella.52 In SSCP, heat-denaturation is used to induce strand separation in PCR-amplified products, which are then subjected to nondenaturing gel electrophoresis.

855

During electrophoresis, even minor conformational changes induced by innate point mutations in the DNA strands can alter gel migration patterns of individual DNA strands and present a means by which genotypic markers can be generated and evaluated. The results indicate that a high level of sequence variation exist in the expansion segment V (ESV) region of the large subunit ribosomal DNA (lsrDNA)63 and in the D3 domain of nuclear ribosomal DNA. Based on these two markers, all of the recognized species and genotypes can be distinguished, with the exception of the genotype T12 which has not been tested. The results also show substantial sequence variation among isolates of the same taxon. Thus, if used to analyze a variable region of DNA such as the Trichinella ESV sequences, SSCP shows great promise for defining both interand intraspecific variation. PCR-restriction fragment length polymorphism. For reasons described above,62 researchers continue to evaluate sequence variation within distinct genes to develop speciesspecific PCR primers or to generate unique restriction enzyme  banding patterns from PCR-amplified gene fragments using genus-specific primers.55,57 Degenerate primers based on the CO I gene of mitochondrial DNA (mtDNA), have been used to amplify and sequence DNA from nine Trichinella taxa and subsequently differentiated them by RFLP analysis.55 Multiplex-PCR. To unequivocally distinguish all currently recognized Trichinella species and the T6 genotype, a multiplex PCR test that uses five primer sets based on the ESV sequences and the internal transcribed spacer (ITS) regions 1 and 2 has been developed.59 The multiplex PCR eliminates the need to perform multiple PCR reactions. When the five primer sets are combined into a single mix and PCR is performed on individual parasites, a distinct and simple agarose gel profile is generated for each of the eight species and the T6 genotype. Each taxon is characterized by no more than two PCR fragments. The ability to perform this test on individual larvae makes it ideal for samples where often only a few larvae are available. The technology as originally described, used nested PCR for the individual larval analysis;59 however, the protocol has been improved and a single round PCR has been generated to work on single larvae.60 The molecular identification of single larvae is carried out with a multiplex-PCR analysis, which allows for the identification of eight species (T. spiralis, T. nativa, T. britovi, T. pseudospiralis, T. murrelli, T. nelsoni, T. papuae, and T. zimbabwensis), the T6 genotype, and three populations of T. pseudospiralis (from the Australian, Nearctic, and Palearctic regions). Since the genotypes T8 and T9 show the same band pattern as T. britovi by multiplex PCR, a PCR-RFLP analysis of the gene encoding for the CO I protein can be used to distinguish between T. britovi and genotype T9.55 To distinguish between T. britovi and genotype T8, a standard PCR based on the ITS2 sequence can be used.24 Reverse line blot hybridization (RLB). An RLB assay has been developed to identify different Trichinella genotypes.61 The RLB assay can be used to detect and identify

856

the most of Trichinella taxa (T. spiralis, T. britovi, T. nativa, T. murrelli, T. nelsoni, and T. pseudospiralis, and Trichinella T6 and T8); it is based on the hybridization of the amplified 5S ribosomal DNA intergenic spacer regions to specific, membrane-bound oligonucleotide probes.

63.2  Methods 63.2.1 Sample Collection and Preparation 63.2.1.1 Reference Material A major resource of vital parasite material and information is the International Trichinella Reference Centre (ITRC) in Rome, Italy (www.iss.it/site/Trichinella/index.asp), which was established in 1990 and whose biobank currently contains more than 2,000 isolates from around the world. For each isolate, the ITRC database includes information on the results of molecular typing, the host species, and the geographical origin, as well as other epidemiological information.64 63.2.1.2 Hosts and Preferential Muscles The best candidates for detecting Trichinella are large carnivorous and omnivorous animals (mammals, birds, and reptiles) that engage in scavenger and cannibalistic behavior and are at the top of the food chain. The most common candidates in Nearctic and Palearctic regions are the polar bear, brown bear, black bear, wolf, red fox, lynx, bob cat, and racoon dog; in other regions, some examples of the most common candidates are viverrides, spotted hyena and lion in the Ethiopian region and carnivorous marsupials in Tasmania. At the farm level (i.e., in the domestic habitat), the best candidate is the domestic pig; other candidates include synanthropic animals such as rats, foxes, mustelids, armadillos, domestic cats and dogs, and farmed crocodiles. Trichinella infection has also been detected, though rarely, in horses and sheep, which were the source of human infection, yet infected herbivorous animals represent an offshoot of the infection occurring in pigs.1 As mentioned, Trichinella is most commonly identified based on the analysis of muscle larvae, which are very easy to collect from both animals and humans. Although PCRderived methods have been used to perform molecular diagnosis of Trichinella infection by detecting DNA directly in the muscle (i.e., without performing artificial digestion to collect the larvae), these methods have been shown to have a low sensitivity because of the huge amount of muscle DNA, which dilutes the larva target sequence DNA. These methods are also unfeasible because of the great amount of Proteinase K that would be needed to digest the muscle sample. Muscle samples should be collected from the preferential muscles (i.e., those with the highest density of larvae), which vary by host species. However, as a general rule, the base of the tongue can be considered as the preferential muscle, though the superficial layer should be removed before sample preparation, since it cannot be digested and prevent to read the sediment. Other important muscles are: the pillar of the diaphragm and the masseter in swine and rodents; the

Molecular Detection of Foodborne Pathogens

anterior tibial in carnivorous mammals; and the masseter in carnivores, horses, crocodiles, monitor lizards. Human biopsies are generally taken from the deltoid muscle. Other important considerations when choosing the specific muscle to be used for digestion, are the digestibility of the muscle, which can vary greatly,65 and the amount of muscle, free of fat and fascia, which should be tested. Because the average number of larvae per gram of preferential muscle tissue generally ranges from 0.1 to 1.0, the chance of detecting larvae increases with the amount of tissue sampled. For meat products (e.g., sausage), before digestion the muscle tissue should be separated from the fat and other tissues and then cut into small pieces and rehydrated (if necessary) at + 4°C in a saline solution for 1–2 days. If digestion is not performed immediately, yet within 2  weeks of the animal’s death, then the samples should be stored at + 4°C. If digestion is performed more than 2 weeks after the animal’s death, then the samples should either be frozen at –20°C, without interrupting the cold chain before digestion, or stored at room temperature for 5–6 months in a 0.1% trimerosal (merthiolate) solution. 63.2.1.3  Parasite Isolation and Preservation The magnetic stirrer method is considered the gold standard for isolating larvae because it is the only method that has been subjected to validation studies.66 The equipment listed in this section and in all of the following sections includes that which may not be present in the laboratory (i.e., standard materials such as beakers and pipettes are not always listed). Reagents: (i) 37% hydrochloric acid; and (ii) pepsin, strength: 1:10000 NF (US National Formulary). Equipment: (i) A glass beaker with a capacity of at least 3 l; (ii) a thermometer that can measure temperatures up to 60°C; (iii) a magnetic stirrer with a thermostatically controlled heating plate and a Teflon-coated stirring rod; (iv) a professional blender with a removable blade and a glass pitcher with a capacity of at least 500 ml; (v) a conical glass separatory funnel, with a capacity of at least 2 l, preferably fitted with a Teflon safety plug; (vi) a sieve with a mesh size of 180 µm and a diameter of 11 cm; (vii) a glass funnel whose diameter is large enough so that it can contain the sieve; (viii) a gridded Petri dish; and (ix) a stereo-microscope (15–40 × magnification), with a substage transmitted light source of adjustable intensity. Procedure: A maximum of 100 g of muscle tissue from the preferential muscle should be tested. The sample should be taken near the muscle insertion, where the greatest concentration of larvae is located. The ratio of muscle (weight) to digestion fluid (volume) should be 1:20. The quantities reported below are for digesting 100 g of muscle tissue. If less than 100 g of meat is used, the amount of digestion fluid should be proportionally reduced. (1) In a beaker, add 16 ml of hydrochloric acid to 2.0 l of tap water that has been preheated to 44–45°C. (2) Place the beaker on the preheated plate of the magnetic stirrer and stir with the Teflon stirring rod.

Trichinella

(3) Add 10 g of pepsin. (4) Cut the muscle sample into small pieces (0.5–2 g each), removing all non-muscle tissue (tendons, fat, etc.). (5) Place the pieces of muscle in a professional blender and mince. (6) Transfer the minced meat to the beaker containing the water, pepsin, and hydrochloric acid, leaving the beaker on the heated plate; to remove any remaining meat from the blender, remove the blade and immerse it in the beaker containing the digestion fluid; rinse the blender glass pitcher with a small quantity of digestion fluid and pour into the beaker. (7) Cover the beaker with aluminum foil. The magnetic stirrer must be adjusted so that the digestion fluid maintains a constant temperature of 38–40°C throughout the procedure. The digestion fluid must be stirred at a sufficiently high speed to create a deep whirl without splashing (during this process, the muscles are digested and the larvae remain in the fluid). The digestion time is generally around 30  min, though more time may be needed (not to exceed 60 min) for certain types of meat (e.g., tongue, game meat). The process is complete when undigested muscle is no longer present in the fluid. (8) Place the sieve in the glass funnel, which has been placed in the separatory funnel. (9) Pour the digestion fluid through the sieve (which retains any undigested tissues but allows the passage of the larvae). The digestion process can be considered as successful if no more than 5% of the initial sample (in terms of weight) remains on the sieve; allow the larvae to sediment for 30 min. (10) Open the safety plug of the separatory funnel and release 40 ml of the sediment into a 50 ml tube. (11) Allow the larvae in the separatory funnel to sediment for another 10 min, to clarify the suspension. (12) Discard 30 ml of supernatant and pour the remaining 10 ml of sediment into a gridded Petri dish. (13) Rinse the 50 ml tube with 10 ml of tap-water, shake, and add to the sediment in the gridded Petri dish. (14) Examine the sample in the Petri dish by a stereo-microscope (15–40 × magnification) for the presence of Trichinella larvae, as soon as possible after sedimentation. Larvae should then be stored in absolute ethanol, which should be done immediately if the larvae are dead (C-shaped or comma shaped), to avoid DNA destruction. 63.2.1.4 DNA Purification from Single Larvae Reagents: (i) DNA IQ™ System kit, code DC6701 (Promega, Madison, WI); and (ii) tissue and hair extraction kit, code DC6740 (Promega, Madison, WI). Incubation buffer (IB) and lysis buffer (LB) are provided with the two commercial kits, yet before use they should be supplemented with Proteinase K and DTT, according to the manufacturer’s instructions (the supplemented buffers are referred to as “IB + ” and “LB + ”).

857

Equipment: (i) A Petri dish; (ii) a thermoblock with vibration function (e.g., Thermomixer comfort, Eppendorf, USA) and temperature range of 25–90°C; (iii) a magnetic separation stand; (iv) a stereo microscope (15–40 × magnification); (v) 1.5 ml conical tube. Procedure: This procedure is for larvae that have been isolated according to the method described in Section 63.2.1.3. The procedure recommended for the kit is slightly modified for use with single Trichinella larvae. (1) Transfer the ethanol containing the larvae into a Petri dish. (2) Under the microscope, collect single larvae using a 2 µl tip and place one larva each in a 1.5 ml conical tube. (3) Spin the tubes at maximum speed for a few seconds and then make sure that the larva is on the bottom of the tube, using the microscope. (4) Allow the ethanol to evaporate completely, leaving the tube open for a few minutes. (5) Prepare a quantity of IB + and LB + that is sufficient for the number of samples to be analyzed. (6) Add 20 µl of IB + ; (7) incubate at 55°C for 30 min in the thermoblock, shaking at 1,400 vibrations/min. (8) Spin the tubes. (9) Add 40 µl of LB + and mix by pipetting a few times. (10) Add 4 µl of the paramagnetic resin (Tissue and Hair Extraction Kit) and resuspend it by slow vortexing, avoiding the creation of bubbles. (11) Incubate for 5 min at 25°C in the thermoblock, shaking at 1,400 vibrations/min. (12) Place the tubes in the magnetic separation stand and wait for 60 sec. (13) Discard the liquid phase by aspiration, avoiding the dislodging of the resin particles. (14) Add 100 µl of LB + and resuspend the resin particles by vortexing. (15) Repeat steps 12 and 13. (16) Add 100 µl of 1 × wash buffer (Tissue and Hair Extraction Kit) and resuspend the resin particles by vortexing. (17) Repeat steps 12 and 13. (18) Repeat two times steps 16, 12, and 13. (19) Leave the tubes open for 15–20 min to allow the resin particles to dry. (20) Add 50 µl of the eluting buffer (Tissue and Hair Extraction Kit) and gently resuspend the resin particles, without vortexing. (21) Incubate at 65°C for 5 min and shake at 1,400 vibrations/min. (22) Repeat step 12. (23) Collect the liquid phase (about 40–50 µl) and transfer it to a 1.5 ml conical tube. The DNA cannot be checked using a gel because the total amount is very low. (24) Store the DNA samples frozen (–20°C).

858

Molecular Detection of Foodborne Pathogens

63.2.2  Detection Procedures 63.2.2.1  Multiplex PCR Reagents: (i) 2 × PCR Master Mix commercial solution (standard composition: 400 µM dATP, 400 µM dCTP, 400 µM dGTP, 400 µM dTTP, 3 mM MgCl2, 50 U/ml Taq DNA polymerase; (ii) oligonucleotides: (a) ESV target locus, 5′-GTTCCATGTGAACAGCAGT-3′, 5′-CGAAAACATACGACAACTGC-3′; (b) ITS1 target locus, 5′-GCTACATCCTTTTGATCTGTT-3′, 5′-AGACACAATA TCAACCACAGTACA-3′, and 5′-GCGGAAGGATCATTAT CGTGTA-3′, 5′-TGGATTACAAAGAAAACCATCACT-3′; and (c) ITS2 target locus, 5′-GTGAGCGTAATAAAGG TGCAG-3′, 5′-TTCATCACACATCTTCCACTA-3′, and 5 ′ - C A A T T G A A A A C C G C T T A G C G T G T T T- 3 ′, 5′-TGATCTGAGGTCGACATTTCC-3′. These lyophili­zed products are reconstituted with 0.1 × TE at a concentration of 100 pmol/µl; (ii) SetB, the oligonucleotide mixture used for the multiplex-PCR. The mixture is obtained combining an equal volume of the ten oligonucleotides (final concentration of 10 pmol/µl); (iv) genomic DNA (10 ng/µl) purified from a pool of larvae of a Trichinella reference strain (ITRC, Rome, Italy), considered as positive control DNA; and (v) purified DNA of the larvae to be tested (see Section 63.2.1.4). Equipment: A thermocycler. Procedure: The total volume of the amplification reaction is 30 µl. Before the procedure, thaw the following on ice: DNA samples purified from single larvae, 2 × PCR Master Mix, SetB, and positive control DNA. (1) Prepare the amplification solution by adding sequentially: 15 µl 2 × PCR Master Mix, 4 µl water, 1 µl SetB, and 10 µl of DNA purified from single larvae to be tested (see Section 63.2.1.3) or control DNA. When working with multiple samples, a quantity of amplification solution that is sufficient for all of the samples can be prepared all at once; aliquot the solu-





tion in the PCR tubes containing the DNA samples to be tested. (2) Close the tubes, mix by vortexing, and centrifuge at maximum speed for a few seconds. (3) Perform amplification: one cycle of 95°C for 4 min; 35 cycles of 95°C for 20 sec, 55°C for 30 sec and 72°C for 30 sec; one cycle of 72°C for 3 min. It is recommended that a hot start Taq DNA polymerase be used, instead of standard enzymes. (4) At the end of the amplification, centrifuge the tubes at maximum speed for a few seconds. (5) Add the DNA loading buffer and load the samples onto a 2% agarose gel to view the result. (6) Evaluate the size of the amplification bands by comparison with the molecular weight markers. The species is identified by comparing the size of the band(s) shown by the sample with those in Table 63.2 and Figure 63.1.

63.2.2.2 PCR-RFLP to Distinguish between Trichinella britovi and Trichinella T9 When using multiplex-PCR, larvae belonging to the T9 genotype display the same band pattern as T. britovi larvae. To distinguish between these two taxa, a PCR-RFLP method based on the CO I mitochondrial gene sequence has been developed.55 According to the available information, the T9 genotype is circulating only in Japan; for this reason, it is suggested that this method be used only if the larvae with a T. britovi band pattern by multiplex-PCR originated from this country. Reagents: (i) 2 × PCR Master Mix; (ii) oligonucleotides for CO I, as target locus: L6625, F 5′-TTYTGGTT YTTCGGKCACCC-3′; H7005, R 5′-ACGACGTAGTAGGTR TCRTG-3′; (iii) reference DNA from T. britovi and Trichinella T9 (ITRC, Rome, Italy); and (iv) purified DNA of the larvae to be tested (see Section 63.2.1.4). Equipment: A thermocycler.

Table 63.2 Dimension of the Expected Amplification Products (in Base Pairs) for Each Taxon by Multiplex PCR T. T. T. pseudospiralis pseudospiralis pseudospiralis T. T. T. Palearctic Nearctic Tasmanian spiralis nativa* britovi** Isolates Isolates Isolates Genotype

T1

ESV ITS1 ITS2

173

T2, T12 127

T3, T8, T9 127 253

T. Trichinella murrelli T6

T. nelsoni

T. T. papuae zimbabwensis

T4A

T4B

T4C

T5

T6

T7

T10

T11

310

340

360

127

127 210

155

240

264

316

* Not only Trichinella nativa but also Trichinella T12 display a single band of 127 bp. ** Not only Trichinella britovi, but also Trichinella T8 and Trichinella T9 display two bands of 127 bp and 253 bp.

404

859

Trichinella

Ts Tna Tb

TpP TpN TpA Tm T6 Tne Tp

Tz

450 350 250

450 350 250

150

150

Figure 63.1  Photograph of an ethidium bromide-stained 2.5% agarose gel under ultraviolet light illumination showing the multiplex PCR amplification of single larvae of 11 taxa of Trichinella. The samples are as follows: Marker (ladder 50), sizes are in base pairs; Ts, T. spiralis; Tna, T. nativa; Tb, T. britovi; TpP, T. pseudospiralis (Palearctic isolate); TpN, T. pseudospiralis (Nearctic isolate); TpA, T. pseudospiralis (Tasmanian isolate); Tm, T. murrelli; T6, Trichinella T6; Tne, T. nelsoni; Tp, T. papuae; and Tz, T. zimbabwensis. The genotypes Trichinella T8 and Trichinella T9 show the same band pattern of T. britovi. The genotype Trichinella T12 shows the same band pattern of T. nativa.

Procedures: The procedure consists of two steps: (i) PCR amplification of samples; and (ii) restriction analysis of the PCR products. (i) PCR amplification: The total volume of the amplification reaction is of 50 µl. (1) Prepare the amplification solution by adding sequentially: 25 µl 2 × PCR Master Mix, 13 µl water, 1 µl of each primer, and 10 µl of DNA extract from the single larvae to be tested (see Section 63.2.1.3). (2) Apply the following cycling program: one cycle of 94°C for 4 min; 35 cycles of 94°C for one min, 48°C for 1 min, and 72°C for 1 min; 1 cycle of 72°C for 3 min. It is recommended that a hot start Taq DNA polymerase be used instead of standard enzymes. (3) At the end of the amplification, centrifuge the tubes at maximum speed for a few seconds. (4) Add the DNA loading buffer and load the samples onto a 2% agarose gel to view the result (both T. britovi and Trichinella T9 display a 419 bp band). The amount of DNA product present in each sample should be evaluated by eye. (5) Store the amplified products at –20°C until use. (ii) Restriction analysis: Set the reactions keeping the tube on ice. (1) For each sample, transfer at least 100 ng of the amplified product into a 1.5 ml reaction tube. (2) Set the volume to 10 µl with water. (3) Add to the reaction tube: 5 µl of 10 × MseI reaction buffer, 1 µl of MseI (10 U/µl) restriction enzyme, and water up to 50 µl. When working with multiple samples, a quantity of amplification solution that is sufficient for all of the samples can be prepared all at once; aliquot the solution in 1.5 ml tubes containing the PCR products to be tested. (4) Mix and incubate at 37°C for 60 min. (5) Stop the reaction by heating at 65°C for 20 min and then spinning for a few seconds. (6) Visualize the results by 2% agarose gel electro­ phoresis.

(7) Evaluate the results according to the expected size pattern: for T. britovi five bands of 22, 62, 64, 70, and 201 bp; for Trichinella T9, two bands of 92 and 327 bp (Table 63.1). 63.2.2.3 PCR to Distinguish between Trichinella britovi and Trichinella T8 By multiplex-PCR, the larvae of Trichinella T8 display the same band pattern as T. britovi larvae. To distinguish between these two taxa, a PCR method based on a 21 bp deletion in the ITS2 sequence of Trichinella T8 has been developed.24 According to the available information, the T8 genotype is circulating only in sub-Saharan Africa; it is thus recommended that this method only be used if the larvae with a T. britovi band pattern by multiplex-PCR originate from this region. Reagents: (i) oligonucleotides: ITS2, as target: ITS2G.F 5′-CCGGTGAGCGTAATAAAG-3′, ITS2G.R, 5′-TACACACAACGCAACGAT-3′;24 (ii) reference DNA from T. britovi and from Trichinella T8 (ITRC, Rome, Italy); and (iii) purified DNA of Trichinella larvae to be tested (see Section 63.2.1.4). Equipment: A thermocycler. Procedure: The total volume of the amplification reaction should be of 30 µl. (1) Prepare the amplification solution by adding sequentially: 15 µl 2 × PCR Master Mix, 3 µl water, 1 µl of each primer and 10 µl of DNA extract from single larvae or control DNA. When working with multiple samples, a quantity of amplification solution that is sufficient for all of the samples can be prepared all at once; aliquot the solution in the PCR tubes containing the DNA samples to be tested. (2) Close the tubes, mix by a vortex, and centrifuge at maximum speed for a few seconds. (3) Use the following cycling program: one cycle of 94°C for 4 min; 35 cycles of 94°C for 30 sec, 51°C for 30 sec, and 72°C for 1 min; one cycle of 72°C for 3 min. It is recommended that a hot start Taq DNA polymerase be used instead of standard enzymes.

860

Molecular Detection of Foodborne Pathogens

M

1

2

3

4

5

6

7

8

9

M

150 100 50

Figure 63.2  Photograph of an ethidium bromide-stained 2.5% agarose gel under ultraviolet light illumination showing the PCR amplification of single larvae of Trichinella britovi and Trichinella T8. The samples are as follows: M, (ladder 50), sizes are in base pairs; lines 1–3 and 8, Trichinella T8; lines 4–7, T. britovi; line 9, negative control.

(4) At the end of the amplification, centrifuge the tubes at maximum speed for a few seconds. (5) Add the DNA loading buffer, and visualize the results by 2% agarose gel electrophoresis. (6) Evaluate the result according to the expected size pattern: T. britovi, band of 125 bp; Trichinella T8, band of 104 bp (Figure 63.2). 63.2.2.4 SSCP A simple, nonisotopic (“cold”) SSCP approach for the analysis of genetic variation within and among the currently recognized taxa of the Trichinella genus using the variable ES5 as the target region has been developed.52 Reagents: (i) 2 × PCR master mix solution (400 µM dATP, 400 µM dCTP, 400 µM dGTP, 400 µM dTTP, 3 mM MgCl2, 50 U/ml Taq DNA polymerase, Promega code M7501); (ii) denaturing DNA loading buffer (10 mM NaOH, 95% formamide, 0.05% of both bromophenol blue and xylene cyanole); (iii) GMATM S-50 precast gels; (iv) oligonucleotides: ESV target locus: oTsr4 5′-GTTCCATGTGAACAGCAGT-3′ and oTsr3 5′-GCGATTGAGTTGAACGC-3′;63 D3 domain target locus: D328-26 5′-ACCCGTCTTGAAACACGGA-3′ and D328-25R 5′-GATTAGTCTTTCGCCCCTA-3′; (v) genomic purified DNA (10 ng/µl) from a pool of Trichinella larvae of each taxon (ITRC, Rome, Italy) Equipment: (i) A thermocycler; and (ii) horizontal electrophoresis apparatus connected to a cooling system. Procedure: The total volume of the amplification reaction should be of 50 µl. (1) Prepare the amplification solution by adding sequentially: 25 µl 2 × PCR Master Mix, 13 µl water, 1.0 µl of each primer (25 pmol), and 10 µl of DNA extract from single larvae or control DNA. When working with multiple samples, a quantity of amplification solution that is sufficient for all of the samples can be prepared all at once; aliquot the solution in the PCR tubes containing the DNA samples to be tested. (2) Close the tubes, mix by vortexing and centrifuge at maximum speed for a few seconds. (3) Apply the following cycling program: one cycle of 95°C for 5 min; 35 cycles of 94°C for 30 sec, 55°C





for 30 sec, and 72°C for 30 sec; one cycle of 72°C for 5 min. It is recommended that hot start Taq DNA polymerase be used instead of standard enzymes. (4) At the end of the amplification, centrifuge the tubes at maximum speed for a few seconds. (5) Add 1 volume of denaturing DNA loading buffer into each sample tube. (6) Load the samples onto a 2% agarose gel to check the amplification products. Evaluate by eye the amount of DNA product in each sample, and evaluate the volume to submit to the SSCP run to have signals of similar intensity. (7) Denaturate the samples at 94°C for 15 min. (8) Perform snap-cooling on individual samples (12 ml) on a freezer block (–20°C). (9) Load samples onto precast GMATM S-50 gels (96 × 261 mm). (10) Run electrophoresis for 14 h at 72 V and constant temperature of 7.2°C in a horizontal apparatus connected to a cooling system. (11) Stain the gel with ethidium bromide (0.5 μg/ml) for 15 min. (12) Destain the gel in water for 15 min and take a photograph of the results. (13) For the identification of the taxa, compare the band patterns of the tested samples with those of the reference DNAs.

63.3 Conclusions and Future Perspectives The molecular methods used to identify parasites of the genus Trichinella have been developed with muscle larvae, which can be easily collected from infected animals, meat samples, and human biopsies. Attempts to detect migrating newborn larvae in blood or buffy coat samples, using a PCR-derived method, have failed because the newborn larva are produced intermittently, making it difficult to identify the period of larval production by females parasitizing the gut. This problem could potentially be resolved by improving methods for extracting DNA from blood, which would allow the infection to be diagnosed earlier. With regard to the possibility of using a PCR-derived method directly on muscle samples (i.e., without first performing artificial digestion), this is not feasible, given that, on average, there are only one to ten larvae (or less) per gram of muscle, so that the amount of larva DNA would be greatly exceeded by the amount of host DNA and thus not detectable. Only if the quantity of muscle larvae were exceptionally high (e.g., 100 larvae/g or more) would the primers succeed in recognizing the target DNA. Thus PCRderived methods cannot be used to diagnose Trichinella infection but only to identify the parasite at the species or genotype level. A possible goal for the future would be to develop new PCR-derived methods that would allow a diagnosis to be made on muscle samples with a low worm burden.

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One of the advantages of PCR-derived methods is that they are sensitive enough to identify the species or genotype based on the analysis of a single larva or fragment of larva. This is extremely important for detecting mixed infections, which can provide important information on host-parasite contacts and on the role of immunity in preventing the development of a secondary infection.1 Since the discovery that the genus Trichinella is characterized by more than one species, scientists have developed new and simple methods for identifying the various taxa. At present, PCR-derived methods have proven to have both the highest sensitivity and the highest specificity. However, when using these methods, it should be kept in mind that the ability to differentiate isolates should not be equated with their classification as a unique taxon. As mentioned, some PCR-derived methods are so sensitive that the species or genotype of the parasite can be identified based on only 1/15 of larval DNA. This amount of DNA could probably be further reduced using new technologies such as real-time PCR, although the cost-benefit ratio of applying such methods would obviously be quite high. Anomalies can occur when analyzing individual larvae by multiplex PCR. Genotype-specific bands generated from the ITS primer pairs are occasionally absent from the band pattern. This is most likely the result of a poor amplification of the ITS region, together with poor quality genomic DNA extracted from individual larvae. Alternatively, absolute sequence conservation may not exist among individual parasites within a given population at the genotype-specific primer binding sites in the multiplex PCR. Less than 100% sequence similarity at the 3′-end could also cause periodic loss of bands. Since Trichinella parasites reach humans through the consumption of meat, it would be very useful to identify microsatellite loci which could be used as isolate markers to trace the source of infection. This would contribute to the control and eradication of foci in the domestic habitat and to identifying the region of origin of infected wild animals, so as to inform local hunters of the risk of zoonotic transmission constituted by certain practices, such as that of leaving carcasses in the field after skinning, or discarding entrails.1 Tracing the source of infection has become increasingly important with the expansion of the global market of food products. For example, in a large outbreak of trichinellosis for the consumption of horse meat in France, the identification of the etiological agent as T. murrelli allowed the origin of infection to be traced back to the USA, given that this species only exists in North America.

















References

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Molecular Detection of Foodborne Pathogens 43. Dupouy-Camet, J. and Bruschi, F. Management and diagnosis of human trichinellosis. In FAO/WHO/OIE Guidelines for the Surveillance, Management, Prevention and Control of Trichinellosis. Dupouy-Camet, J. and Murrell, K.D. (Eds.). World Organisation for Animal Health, Paris, 2007. 44. Kociecka, W. Trichinellosis: human disease, diagnosis and treatment. Vet. Parasitol., 93, 365, 2000. 45. Capó, V.A., Despommier, D.D. and Polvere, R.I. Trichinella spiralis: vascular endothelial growth factor is up-regulated within the nurse cell during the early phase of its formation. J. Parasitol., 84, 209, 1998. 46. Dick, T.A. et al. The use of the polymerase chain reaction to identify porcine isolates of Trichinella. J Parasitol., 78, 145, 1992. 47. Soulé, C. et al. Differentiation of Trichinella isolates by polymerase chain reaction. Parasitol. Res., 79, 461, 1993. 48. Bandi, C. et al. Random amplified polymorphic DNA technique for the identification of Trichinella species. Parasitology, 107, 419, 1993. 49. Bandi, C. et al. Random amplified polymorphic DNA fingerprints of the eight taxa of Trichinella and their comparison with allozyme analysis. Parasitology, 110, 401, 1995. 50. Wang, H. et al. Restriction fragment length polymorphism (RFLP) analysis of genomic DNA of 5 strains of Trichinella spiralis in China. Chin. Med. Sci. J., 10, 131, 1995. 51. Gasser, R.B. et al. PCR-SSCP of rDNA for the identification of Trichinella isolates from mainland China. Mol. Cel. Probes, 12, 27, 1998. 52. Gasser, R.B. et al. Nonisotopic single-strand conformation polymorphism analysis of sequence variability in ribosomal DNA expansion segments within the genus Trichinella (Nematoda: Adenophorea). Electrophoresis, 25, 3357, 2004. 53. Wu, Z. et al. Polymerase chain reaction primers to identify Trichinella spiralis or T. pseudospiralis. Parasitol. Int., 46, 149, 1997. 54. Wu, Z., Nagano, I. and Takahashi, Y. The detection of Trichinella with polymerase chain reaction (PCR) primers constructed using sequences of random amplified polymorphic DNA (RAPD) or sequences of complementary DNA encoding excretory-secretory (E-S) glycoproteins. Parasitology, 117, 173, 1998. 55. Nagano, I. et al. Identification of Trichinella isolates by polymerase chain reaction-restriction fragment length polymorphism of the mitochondrial cytochrome c-oxidase subunit I gene. Int. J. Parasitol., 29, 1113, 1999. 56. Appleyard, G.D. et al. Differentiation of Trichinella genotypes by polymerase chain reaction using sequence-specific primers. J. Parasitol., 85, 556, 1999. 57. Wu, Z. et al. Polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP) for the identification of Trichinella isolates. Parasitology, 118, 211, 1999. 58. Wu, Z. et al. DNA fingerprints of Trichinella as revealed by restriction fragment length polymorphism and single-strand conformational polymorphism (RFLP-SSCP). Mol. Cell. Probes, 14, 291, 2000. 59. Zarlenga, D.S. et al. A single multiplex PCR for unequivocal differentiation of six distinct genotypes of Trichinella and three geographical genotypes of Trichinella pseudospiralis. Int. J. Parasitol., 29, 1859, 1999. 60. Pozio, E. and La Rosa, G. PCR-derived methods for the identification of Trichinella parasites from animal and human samples. Methods Mol. Biol., 216, 299, 2003.

Trichinella 61. Rombout, Y.B., Bosch. S. and Van Der Giessen, J.W. Detection and identification of eight Trichinella genotypes by reverse line blot hybridization. J. Clin. Microbiol., 39, 642, 2001. 62. MacPherson, J.M. et al. Variability of the random amplified polymorphic DNA assay among thermal cyclers, and effects of primer and DNA concentration. Mol. Cell. Probes, 7, 293, 1993. 63. Zarlenga, D.S. and Dame, J.B. The identification and characterization of a break within the large subunit ribosomal RNA of Trichinella spiralis: comparison of gap sequences within the genus. Mol. Biochem. Parasitol., 51, 281, 1992.

863 64. Pozio, E. et al. Twelve years of activity of the International Trichinella Reference Centre. Parasite, 8, S44, 2001. 65. Kapel, C.M., Webster, P. and Gamble, H.R. Muscle distribution of sylvatic and domestic Trichinella larvae in production animals and wildlife. Vet. Parasitol., 132, 101, 2005. 66. Nöckler, K. and Kapel, C.M.O. Detection and surveillance for Trichinella meat inspection and hygiene, and legislation. In FAO/WHO/OIE Guidelines for the Surveillance, Management, Prevention and Control of Trichinellosis. Dupouy-Camet, J. and Murrell, K.D. (Eds.). World Organisation for Animal Health, Paris, 2007

Index A Acanthamoeba spp. Acanthamoeba keratitis (AK) diagnosis, 640–641 DNA extraction methods, 641 infection and clinical signs, 640 PCR methods, 641 topical treatment, 640 conventional diagnosis antibody testing, 642 culture, 642 histopathologic examination, 642 infections, 640–642 life cycle, stages in cyst, 640 trophozoite, 639 methods and procedure, 646–647 DNA extraction from corneal and noncorneal samples, 643 genotyping based on sequencing analysis, 645–646 reagents and equipment, 643 real-time PCR assay, 644–645 sample collection, 643 standard PCR detection, 644 molecular diagnosis DNA extraction methods, 642 molecular typing, 642 PCR methods, 642 taxonomy of, 639 Actin as targets, 12; see also Targets for detection techniques Acute viral hepatitis, 71 Adenoviruses classification and morphology, 23–24 detection in foods concentration by PEG precipitation, 28–29 extraction procedure, 28 guanidine thiocianated extraction method, 29–30 materials, 28 quality control, 30 sample preparation, 28 identification and diagnosis antibody-based techniques, 27 cell culture and PCR, combination of, 27–28 culture-based techniques, 27 nucleic acid probes, 27 PCR techniques, 27 infection epidemic keratoconjunctivitis (EKC), 25 follicular conjunctivitis, 25 gastroenteritis, 25 immunocompromised, impact on, 25 obesity, 25 pharyngoconjunctival fever (PCF), 25 respiratory illnesses, 25 routes, 24 water-and foodborne outbreaks, 25–26 occurrences in shellfish, 26 water, 26 properties of, 24 survival in environment and shellfish, 26 Adhesins role in aeromonad virulence, 278 Aerokey II system, biochemical identification of Aeromonas spp., 279

Aeromonas spp. Aeromonadaceae family, 273 Aeromonas lipopolysaccharide, surface antigen (O), 274 A. hydrophila, First and Second Contaminant Candidate List (CCL1 and CCL2), 273 biochemical identification Aerokey II system, 279 detection techniques, 281–282 extracellular hydrolytic enzymes, 274 extra-intestinal infections, 276 genomospecies and phenospecies, 275 infections, 273–274 clinical symptoms, 276 isolation of, 278–279 molecular detection and identification, 279–280 motile aeromonads, antibiotic resistance, 274 optimum growth temperature, 275 reagents and equipment, 281 sample collection and preparation aroA gene primers, 281 fragment length polymorphism (RFLP) analysis using HaeII endonuclease, 281 short rigid pili (S/R type) and long wavy flexible pili (L/W type), 277 taxonomy of, 274–275 virulence factors, 274, 276 aerolysin (AerA), 278 Asao toxin, 278 cell-associated, 277 cholera toxin cross-reactive cytolytic enterotoxin, 278 cytotoxin enterotoxin, 278 extracellular, 278 hemolysins, 278 Alternaria spp. characteristic feature of, 521 classification of, 521 differerence with Stemphylium and Ulocladium genera, 521 disorders in humans and animals, 522 growth temperature and media, 522 laboratory techniques for diagnosis of infections conventional techniques, 523 molecular techniques, 523–524 mechanism of action, 522 method of detection DNA isolation, 524 PCR and sequencing analysis, of ITS-5.8S gene region, 524–525 PCR detection and quantification of AM-toxin gene, 525 sample processing, 524 morphological features, 521 mycotoxin produced by, 522 as plant pathogens, 522 species as human pathogens, 522 spores as airborne allergens, 522 spread and effect of infection, 522–523 tenuazonic acid (AT), disease caused by, 522 Alternariosis (alternariatoxicosis), 522 Aminoglycosides S. marcescens resistance mechanisms aminoglycoside n-acetyltransferases (AAC), 464–465 aminoglycoside O-adenylyltransferases (ANT), 465 aminoglycoside O-phosphotransferases (APH), 465 bifunctional aminoglycoside resistance enzyme, 465

865

866 Anisakis spp. A. brevispiculata and A. paggiae distribution of, 759 alignment of nucleotide and DNA sequences, 764–766 allergies, 759 anisakidosis, 757 A. physeteris and A. pegreffii distribution of, 759 A.simplex C distribution of, 759 occupational hypersensitivity, 759 A. ziphidarum and A. typica distribution of, 759 classification of, 758 detection procedures Cox-2 sequences, identification based on, 763 DNA sequencing, 763 ITS restriction patterns, identification based on, 762 PCR amplification, 762 PCR product purification, 762–763 primers, 762 restriction fragment length polymorphisms protocol, 762 diagnosis and treatment, 759 genetic structure analysis, 758 geographic distribution, 758–759 host preference, 758 life cycle, 758 molecular identification, 759 direct DNA sequencing, 760 multilocus allozyme electrophoresis, 760 multiplex and species-specific PCR, 760 PCR-RFLP, 760 reagents and equipment, 761 pathogenesis, 759 sample preparation DNA purification, 761–762 larvae from fish hosts, isolation of, 761 Arcobacter spp. A. butzleri diarrheal infection, 290 A. cryaerophilus subgroup 1 and 2, 296 A. nitrofigilis nitrogen-fixing bacterium, 290 antimicrobial susceptibility, 292 in cattle, 292–293 C. jejuni infection, 290 detection techniques characterization by ERI C-PCR, 300 genus-specific identification by PCR, 299 PCR protocol, 299–300 species identification by PCR-RFL P, 300 species-specific identification by PCR, 299 in food of animal origin, 294–296 in horses, 294 identification and characterization Campylobacter and differentiation between, 296 cell structure, 296 metabolic activity, 296 isolation of, 298 molecular-based characterization AFLP technique, 298 biotyping, phenotyping methods, 298 enterobacterial repetitive intergenic consensus (ERIC) PCR, 298 PFGE and RAPD techniques, 298 molecular identification combined use of PCR with RFLP, 297 genotyping AFLP technique, 297 PCR-hybridization protocol, 297 PCR method, 297

Index phenotypic tests, 296–297 RFLP method, 297 optimal growth, 290 temperature, 296 outbreaks, 291 pathogenicity, 293 pig as reservoir, 293 in poultry, 293–294 sample preparation, 298 DNA extraction protocol, 299 in situ studies of existence of toxigenic and invasive capacities, 291 in veterinary and human public health, 290 virulence factors cytolethal distending toxin (CDT), 291 homologs of fibronectin binding proteins CadF and Cj1349, 291 invasin protein CiaB, 291 phospholipase PldA and TlyA hemolysin, 291 in vivo study with strains, 291 Artemis Disk Surveillance Programme, 551 Aspergillus spp., 529–530 A. aculeatus, 532 A. carbonarius, 532–533, 536–537, 539–540 aflatoxins, and human health, 530 A. flavus, 531, 534, 536–537, 541, 543 A. japonicus, 532 A. niger, 532–533, 536, 539–540 A. ochraceus, 531, 533, 536–537, 539–540 A. parasiticus, 531, 534, 541–542 A. steynii, 533 A. westerdijkiae, 531, 533, 540 classification and taxonomy of, 530–531 division on basis of mycotoxins Aspergillus aflatoxin producing species, 533–534 Aspergillus ochratoxin A producing species, 532–533 foodborne species section Circumdati, 531–532 section Flavi, 531 section Nigri, 532 future research need, 544 laboratory diagnostic techniques for, 534–535 aflatoxin producing fungi, 537–539 ochratoxin A producing species, 535–537 medicinal and industrial use, 529 molecular detection methods, for aflatoxigenic species in corns, 541–542 in figs, 541 in peanut kernel, 542 in wheat, 542–543 molecular detection methods, for ochratoxigenic species in coffee, 539–540 in grapes, 540–541 morphological features, 530–531 mycotoxins produced by, 530 ochratoxin A (OTA), role of, 530 as plant pathogens, 529–530 Astroviruses animal astroviruses, propagation of bovine embryo kidney (BEK) cells, 36 chicken astroviruses, chicken embryo liver (CEL) and chicken hepatocellular carcinoma (LMH) cells, 36 porcine astrovirus, 36 antibody acquisition and prevalence, 38 classification, 35 cytopathic effect (CPE), 35 detection procedures RT-PCR, 40–41 viral RNA extraction, 40 diagnostic methods, 38–40 environmental factors, stability of, 35

867

Index genome arrangement, 34 full-length genomic RNA (gRNA) and sgRNA, 35 HAstV strains isolation, 36 human illness, characteristics of, 36–37 in human, propagation in HEK cells, 35 identification of, 33 image by electron microscopy, 34 immunity for, 37–38 immunocompromised hosts, role in, 37 infections, 36 clinical symptoms associated with, 37 disease, role in, 38 outbreaks for, 38 isolation of, 34 physical and chemical agents, resistance against, 35 sample preparation, 40 mussel sample for RT-PCR detection, 42 for RT-PCR detection, 42 shellfish for RT-PCR detection, 42 sequencing and phylogenetic analysis, 43 star-like morphology, 34 transmission of, 36 treatment and prevention, 40 virus concentration techniques from foods, 41 Avian influenza (AI) viruses and antigen detection, 51–52 classification of, 49 detection procedures H5 HA subtype by qualitative one step real Time RT-PCR, 58–59 one step RT-PCR for detection of H5 subtype, 57 one step RT-PCR for detection of Type A, 56–57 qualitative real time PCR (M gene), Type A, 58 two step RT-PCR for detection of H7 subtype, 57–58 infections caused by, 50, 54 viremia and viral replication, 55 medical importance and zoonotic implication, 50–51 molecular tests for, 53 hemagglutination inhibition test, 52 nucleic acid amplification protocols, 53 PCR-based protocols for detection of, 52–53 RT-PCR based methods, 54 outbreaks in birds, 49 phylogenetic studies, 54 sample collection, 55 sample preparation for PCR testing cloacal and tracheal/oro-pharyngeal swabs, 56 RNA extraction kit, 56 serological methods agar gel immuno-diffusion (AGID) test, 52 surveillance and monitoring programmes, 51 transportation and storage of specimens, 55–56

B Bacillus spp. B. anthracis detection by RSI-PCR, 136–137 etiological agent of anthrax, 129 toxin genes, 131 whole genome SNP analysis, 131 B. cereus abdominal pain and diarrhea, 131 CHO assay, 134 detection by RT-PCR, 138–139 detection techniques by multiplex PCR, 137–138 diarrheal toxins detection, 134 DNA isolation, 136 ecotypes, 130 food and environmental samples, detection of, 134–135 as foodborne pathogen, 129

food poisoning outbreaks, 131–132 G9241 strains, 131 HBL and NHE, tripartite toxins, 132 INRAAF2 strains, 132 isolation and enumeration, 135–136 MLST schemes, 131 murine hybridoma Ped-2E9 cell model, 133 nausea and vomiting, 132 NVH391/98 and NVH883/00 strains, 132 phylogenetic analysis, 132 sample preparation, 135 species discrimination in, 130 strains isolated from periodontitis in humans, 131 toxin detection, 133 B. licheniformis and B. pumilus, heat-stable toxin-producing, 133 B. mojavensis, cytotoxicity of, 133 B. thuringiensis insect pathogen, 129 B. weihenstephanensis genome sequencing, 133 psychrotolerant species, 130 ecology of, 130 Bacteroides spp. B. fragilis biochemical tests, 312 culture, examination of, 312 enterotoxin gene of, 309–310, 314 enterotoxin production, determination of, 312 fecal specimen, collection and transportation of, 311 molecular methods, role of, 310–311 PCR for neuraminidase gene, 314 specimen processing, 312 biological characteristics, 307 CTn9343, map of, 310 detection techniques boiling method, DNA extraction, 313–314 fecal samples, DNA extraction, 314 phenol-chloroform extraction, DNA extraction, 312 epidemiology, 308–309 pathogenesis, 309 PCR reagents for, 311 Bbovine astroviruses (BAstVs) serotypes of astroviruses, 35 Bovine spongiform encephalopathy (BSE), 113 immunohistochemistry of brain tissues from, 118 Brevictenatae, 521 Brucella spp. antigen tests brucellin allergic skin test, 321 buffered plate agglutination test (BPAT), 320 complement fixation test (CFT), 320 enzyme-linked immunosorbent assay (ELISA), 320 fluorescence polarization assay (FPA), 320 Rose Bengal test (RBT), 320 Brucella-specific detection targets, 14 brucellosis, 317 in animals, 318 bacteraemia and, 319 geographic distribution, 319 in human, 319 zoonotic disease, 319 detection techniques multiplex PCR, 323 primers for, 323–324 real-time PCR assays, 324–325 diagnosis isolation and identification, 320 serological tests, 320–321 identification and differentiation of, 323 infection, 319–320 transmission of, 318

868 molecular diagnostic techniques molecular typing methods, 322–323 PCR techniques, 321–322 multiplex PCR cycling program, 324 pathogenicity and, 318 primers for, 323–324 real-time PCR assay targeting, 325–327 sample collection and preparation, 323 serological tests antibody detection, 321 primary binding assays for antibody detection, 321 taxonomy, 317–318 Burkholderia spp. B. mallei, 332 B. pseudomallei, 331 Burkholderia cepacia complex (BCC) infections, 332–333 Burkh olderia gladioli pathovar cocovenenans bacterial isolation, 338 biology, 335–336 conventional diagnosis, 336–337 detection techniques, 338–339 epidemiology, 333–334 molecular diagnosis, 337 multiplex PCR with, 339 pathogenesis, 334–335 propagation, 338 storage, 338 strains of, 339 taxonomy of, 333 toxoflavin extraction, 338 reagents and equipment, 337–338

C Campylobacter spp. animal models, 348 C. jejuni from food, 348 genome of, 346 infection of, 348 life cycle, 346 LOS and flagella, 348 PCR assays, 350 detection techniques duplex real-time PCR amplification for, 355 multiplex PCR, 354–355 and PCR amplification, oligonucleotide primers and probes, 354 flagella and flagellar motility, 347 infection, 346 campylobacteriosis, 345 Guillian–Barré syndrome (GBS), 345 Miller Fisher syndrome (MFS), 345 isolation and detection rapid methods, 349 traditional methods, 348–349 molecular methods DNA-based methods, 349–350 identification of, 351–352 RNA-based methods, 350 whole-genome-based methods, 352 sample preparation and DNA extraction, 353–354 virulence factors adhesion and invasion, 347 chemotaxis and motility, 346–347 cytolethal distending toxin (CDT), 347 glycosylation system, 347 lipooligosaccharide (LOS) and, 347–348 Candida spp., 549–550 C. albicans, 550–552, 557 colonial appearance, 550

Index C. parapsilosis, 552 detection in food culture process, 554–555 PCR technology, use of, 555–557 detection methods and procedure conventional PCR, 559 real-time PCR, 559–560 sample preparation and extraction, 558–559 diagnosis of candidal infections Candida score, calculation of, 553 colonization index, 553 microscopic or culture detection, 553 nonculture diagnostic techniques, 553, 555 PCR protocols suitable for, 554, 556 diseases caused by, 553–554 geographical distribution of species causing bloodstream infections, 551 growth conditions, 550 incidence rate, for infection, 550–551 morphology of, 550 pathogenicity, study on, 552–553 risk factors, for infection, 551–552 ICU residence as, 551 total parental nutrition (TPN) as, 551 species causing invasive candidal infections, 552 Capsid genes for detection of DNA and RNA viruses, 11 Capsules role in aeromonad virulence, 278 carA gene carbamoyl phosphate synthase in Pseudomonas spp. as target, 13 Cereulide by HEp-2 tissue culture assay, 134 Chicken astroviruses (CAstVs) serotypes of astroviruses, 35 Chronic wasting disease (CWD), 113 Clonorchis spp. biology of, 771–772 classification and morphology digenea, 769 clonorchiasis chemotherapy for, 772–773 clinical symptoms of, 772 foodborne, diagnosis of, 773 C. sinensis adult worms surface and sex, 770 appearance and characteristics, 770 body cavity, 771 CS1/CS2 primers, sensitivity of, 774 distribution of, 769 egg production, 778 excretion system and nervous system, 771 infections with, 772 intestine and eggs, 771 life cycle, 769 light micrographs of, 771 size and shape, 770 detection, eggs in human/animal fecal samples cleanup procedure, 775–776 PCR efficacy of, 776 DNA sequences, 776–777 metacercariae in fish, detection of PCR identification for, 775 sample preparation, 774–775 opisthorchiids, eggs of, 777–778 pathogenesis clonorchiasis, clinical symptoms of, 772 prophylaxis, 772 rediae in snails, detection PCR assay, 773–774 sample collection and preparation, 773 Clostridium spp. C. botulinum botulinum neurotoxin (BoNT), 146, 148 genetic typing, 151

869

Index molecular identification, 151 PCR assay for distinguish between group I and group II, 152–153 phenotypic identification, 150 primers for differentiation between group I and group II strains, 151 repetitive extragenic palindromes (REP), 151 standard detection and isolation, 150–151 strains producing toxin, 146 C. difficile proliferation and infection in human colon, 146 toxin A and B, 146 C. perfringens amplified genetic typing methods, 150 antibiotic-associated diarrhea (AAD) and sporadic diarrheas in humans, 146 Clostridium perfringens beta2 toxin (CPB2), 146 Clostridium perfringens enterotoxin (CPE), 146 combination of probe dyes, 149 cpe gene in, 148 food poisoning outbreaks, 148 genetic typing, 150 genome of, 145 molecular identification, 149 multiplex primers for determination of, 147–148 nonamplified genetic typing methods, 150 phenotypic identification, 149 toxin genes, 149 toxinotypes and toxins, 146 toxin producer, 145 type A food poisoning, 146 C. septicum alpha toxin, 148 fatal traumatic and nontraumatic myonecrosis, 146 C. tetani tetanus (lockjaw) in humans and animals, 146 detection techniques multiplex PCR, 152 sample preparation DNA rxtraction, 151–152 COX2 mitochondrial gene as target for detection of S. cerevisiae, 13 cpn60 gene heat shock protein in bacteria as detection target, 13 CpR1 gene from Cryptosporidium target for food detection systems, 14 Creutzfeldt–Jakob disease (CJD), 113 diagnostic criteria for, 116 immunohistochemical staining with monoclonal antibodies in human brain tissue with, 116 Cryptosporidium spp., 651 as cause of diarrhea, 652 mechanism of infection, 652 classification of, 651–652 detection methods, 655 conventional methods, 655 in drinks, 658–659 in fruit and vegetables, 657 in meat, fish, and shellfish, 658 molecular methods, 655–657 detection procedures nested PCR targeting 587-bp 18S rDNA fragment, 660–661 nested PCR targeting 825-bp 18S rDNA fragment, 660 nested PCR targeting GP60 locus, 661 foodborne outbreaks of, 654 food contamination, sources of, 652–654 foodstuffs, as source of oocysts, 652–653 loci used to detect, 656 oocysts, transmission by, 652 reagents and equipment, for extraction and detection of DNA, 659 recognized species of, 652 sample preparation, 659

Cyclospora spp. C. cayetanensis clinical presentation and, 668–669 molecular techniques, 669–670 phenotypic techniques, 669 classification and, 667–668 detection techniques multiplex PCR, 672–673 PCR with F1E-R2B, F3E-R4B primers and, 671–672 real-time PCR, 673 RFLP with AluI and, 672 sample processing fast DNA spin prep kit, 671 feces and foods, 671 FTA filter purification, 671 phenol-chloroform extraction, 671 water, 671 Cytochrome oxidase genes from mt DNA as target for detection of helminths, 13

D Debaryomyces spp., 565 characteristics of, 566 classification of, 565 detection methods and procedures, 569 RAPD-PCR, 571–572 reagents and equipment, 569–570 RFLP analysis of mitochondrial DNA, 570–571 sample preparation, 569–570 D. hansenii, 565–566 excessive growth, effects of, 567 growth in food, 566–567 infections in humans, 567 molecular methods, for identification, 568 methods based on RFLP, 569 nucleic acid hybridization, 568 PCR techniques, 568–569 physiological and morphological identification, 567–568 structure and reproduction, 565–566 Diphyllobothrium spp. biology and pathogenesis, 781–782 classification and morphology, 781 D. dendriticum infections of, 782 D. ditremum infections of, 782 detection procedure, 785 identification, primers used, 786 diagnostic techniques, 782–783 diphyllobothriasis, 781 Diphyllobothriidae family, 781 D. latum infections of, 782 life cycle of, 781 proglottids and plerocercoid, 783 RFLP profiles, 783 DNA extraction from, 785 D. nihonkaiense RFLP profiles, 783 D. vogeli infections of, 782 internal transcribed spacer 1 (ITS1) and COI gene sequences, 784 molecular diagnostic techniques, 783–784 phylogenetic construction, 784 primers for identification, 786 proglottids and plerocercoid of, 783 reagents and equipments, 785 reported from humans and, 782 sample collection, 785 sequences in Genbank, 782

870 E eap gene in Staphylococcus aureus as detection target, 14 Encephalitozoon, 691 Entamoeba spp. E. histolytica, 677–678 ALA, intestinal infection, 684 antibody detection, 681 asymptomatic infection, 680 conventional PCR assays for, 682–684 culture techniques, 681 detection technique, 686–687 extraintestinal disease, 680 infect humans, 678 isoenzyme analysis, 681 life cycle of, 678–679 microscopy, 681 pathogenesis, 679–680 real-time PCR (RT-PCR), 684–685 sample collection and preparation, 686 stool antigen assays, 681–682 symptomatic infection, 680 treatment of, 680–681 Enterobacter spp., 361 detection techniques PCR identification, 365 real-time PCR assay identification, 365 diagnosis techniques conventional procedures, 362–363 epidemiology and pathogenesis, 361 Cronobacter infections, 362 necrotizing enterocolitis (NEC), 362 molecular procedures amplified fragment length polymorphisms (AFLP), 364 BOX-PCR and REP-PCR, 364 multi-locus VNTR analysis (MLVA), 364 PFGE method, 364 random amplification of polymorphic DNA (RAPD), 364 ribotypes, 364 species-specific detection and identification, 363 subtyping, 363 sample preparation, 364–365 Enterococci microorganisms, 157 cell isolation and counting without pre-enrichment, 169 detection techniques data analysis, 174 flowchart of multiplex-PCR, 173 genus-and species-specific multiplex PCR, 173 DNA extraction conventional PCR for, 169–170 from fermented cereal-based food, 172 real-time PCR, 170 enrichment procedure, 169 fermented meat products, 158 kits used for extraction from food matrixes, 170–172 molecular approaches for identification and detection atpA gene sequences, 168 bacterial ATP synthase, 168 culture methods, 158 D-alanine:D-alanine ligase (ddl) gene, 161 elongation factor EF-Tu, 161 fluorescent in situ hybridization (FISH) probes, 159 genus-and species-specific PCR primers, 162–167 heat shock proteins, 168 manganese-dependent superoxide dismutase, 168 microarray technology, 159 molecular diagnostic methods, 159 muramidase, 161 nucleic acid based, 158 PCR assays, 158

Index penicillin-binding proteins (PBPs), 161 probes for, 160–161 real-time quantitative PCR (qPCR), 159 rRNA gene sequences, 159 TaqMan system, 159 tuf gene, 161 viable but nonculturable (VBNC) microorganisms and, 158 sample homogenization, 168–169 vegetables and olives, 158 Enterocytozoon, 691 Enzyme linked immunosorbent assay (ELISA), 8 Epidemic keratoconjunctivitis (EKC), 25; see also Adenoviruses Escherichia spp., 369 detection techniques m-PCR assay, sequences, 382 PCR reaction mixture in, 382–383 E. coli O157:H7 culture and isolation, 371–373 developed m-PCR assay, 376 ELISA, 374–375 enrichment and IMS, 373–374 infections of, 370–371 latex agglutination test (LAT), 373 microbiological and PCR scheme, 372 nucleic acid microarrays, 380–381 occurrence and, 369–370 PCR, 375–376 phenol chloroform isoamylalcohol (PCI) extraction, 376–377 real-time PCR, 377–380 reverse transcriptase (RT)-based assays, 380 Shiga-toxin producing, 369 tiered microbiological and PCR scheme, 372 reagents and equipment, 382 sample preparation, 382 type III secretion system (TTSS), 371 Esculin hydrolysis, biochemical identification of Aeromonas spp., 279 European Food Safety Association (EFSA), 595 Exotic ungulate encephalopathy (EUE), 115

F Fasciola spp. biology, 789–790 detection procedures, 791 fascioliasis, 790 Fasciolidae family, 789 F. gigantica distribution of, 789 life cycle of, 790 F. hepatica distribution of, 789 life cycle of, 789–790 propensity for liver, 789–790 identification and diagnosis differentiation from definitive hosts, 790 infection in snails by PCR, detection of, 790–791 medical importance, 790 pathogenesis, 790 reagents and equipment, 791 sample preparation and DNA isolation, 791 Feline spongiform encephalopathy (FSE), 115 Fluorescence in situ hybridization (FISH) technique Listeria monocytogenes, detection of, 8 Fluoroquinolones S. marcescens resistance to, mechanisms of active efflux pump, 462–463 gyrA, mutations, 462, 464 Qnr determinants, 464 Foodborne illness, 1 pathogen detection techniques

871

Index assay design and data analysis software, 10 combined detection methods, 9 in complex matrices, sample preparation, 3 immunological assays, 8–9 microfabrication and microfluidics, 9 nucleic acid based, 3–6 targets for, 10–14 typing, 9 validation, 14 Food safety foodborne pathogens, detection of, 2 polymerase chain reaction (PCR) inhibitors, 3 Fusarium spp., 577 classification of, 577–578 F. graminearum species complex, 578–579 F. oxysporum species, 579 Gibberella fujikuroi species complex, 578 diagnosis of, 581–582 fumonisins-producing species, 582–583 surface plasmon resonance (SPR) biosensors, 585 trichothecenes producing species, 583–585 future research program, 588 molecular detection of fumonisins-producing fusarium spp. in maize, 585–586 molecular detection of trichothecenes fusarium spp. in wheat, 586–587 quantitative detection of fumonisins-producing fusarium spp. in maize, 586–587 spores types, 578 toxic metabolites by, 579–581 fumonisins, 580 trichothecenes, 579–580

G Gerstmann–Sträussler–Scheinker (GSS), 115 Giardia spp. antigenic variations, 703 classification of, 702 cyst passage rate, 703 detection techniques DNA sequencing, 712 PCR reaction, 712 sequencing analysis, 712–713 diagnosis EnteroTest, 704 G. duodenalis assemblages of, 702 prevalence of, 702 giardiasis, 701 infection outbreaks of, 702–703 life cycle of, 702 molecular methods genotyping tools, 704–708 molecular tools for analysis of environmental samples, 709 PCR assay, 704 reagents and equipment, 710 subtyping tools, 709 in raw vegetables and fruit salad, 702 sample preparation DNA extraction, 711 fecal from, 710 food and water samples, centrifugation of cysts, 711 food and water samples, extraction of cysts, 710–711 transmission, drinking and recreational water, 702 Giardin protein as detection target, 12 Glycerophospholipid cholesterol acyl-transferase (GCAT) role in aeromonad virulence, 278 Granulomatous amoebic encephalitis (GAE), 641–642

H Helicobacter spp. cdt pathogenetic gene, 195 cyclodextrin and culture growth, 182 detection methods for, 182 immunological methods, 183–184 LAMP, 186 molecular techniques, 184 multiplex-PCR, 185 nested PCR, 185 PCR, 184 PCR-DGGE, 186 PCR-RFLP, 185–186 reagents and equipment, 186 real-time PCR, 185 RT-PCR, 185 Southern hybridization, 186 16S rRNA gene, 195 standard PCR, 185 TaqMan-PCR, 185 Taq polymerase, 186–187 target genes, 185 urea breath test (UBT), 184 Helicobacter heilmannii culture study, 195 Helicobacter pylori infection with peptic ulcers and gastric cancer, 188 PCR protocols for detection, 189–194 scanning electron micrograph of, 182 host specificity, 195 infection, 181 isolation, 181 sample collection and preparation biopsies and endoscopy of gastric mucosa, 187 DNA extracted from gastric juice, 187 DNA purification and isolation, 188 feces from, 187 mixed infection with clarithromycin-resistant and-susceptible, 187 plaque from oral cavities, 188 species and hosts, 183 structure, 181 Hepatitis A virus (HAV) classification and genetic diversity 3C/3D region junction, 64 subgenotypes, 64 VP1/P2A region junction, 64 VP3/ VP1 region junction, 64 detection techniques nested PCR for, 69 reverse-transcription PCR (RT-PCR), 68–69 sequence analysis and genotype identification, 69–70 diagnosis molecular techniques, 64 genome, 64 genus Hepatovirus within Picornaviridae family, 63 infection foodborne and waterborne, 64 severe and fulminant hepatic failure (FHF), 63 oligonucleotide primers for detection and sequencing from clinical specimens, 65 specimen collection and RNA extraction, 68 Hepatitis E virus (HEV) classification and genetic diversity serotypes, 66 detection and sequencing from clinical specimens oligonucleotide primers for, 67 reagents and equipments, 67–68 detection techniques nested PCR, 70 RT-PCR, 70

872 sequence analysis and genotype identification, 70–71 genome organization, 66 infection diagnosis, 66 foodborne and waterborne, 66 liver abnormalities, viremia and fecal shedding, 65 molecular techniques, 66 route and mechanism, 66 zoonotic foodborne transmission, 66 member of Hepeviridae family, 66 specimen collection and RNA extraction, 68 Heterophyidae family Centrocestus spp. fish hosts, 796 infection, 797 clinical manifestations clinical symptoms, degree of, 797 heterophyiasis, intestinal histopathology caused, 797 clinical survey, 797 detection procedures endonuclease digestion, 802 fish-infesting trematodes, PCR assays for, 801–802 interpretation of, 802 PCR amplification, 802 diagnosis amplicons, 798 heterophyid trematode sequences, 799–800 heterphyids rDNA, alignments of, 799–800 molecular tools in, 800 PCR amplification, gene targets for, 799 PCR-based methods, 800–801 precise identification of, 798 probes, 798–799 digenea, life cycle of, 798 Haplorchis spp. infection, 797 Heterophyes spp. fishborne zoonoses, 795–796 foodborne trematode infections, 796–797 heterophyid trematodes, 796 human trematodiasis, 795 sample collection and preparation, 801 trematodes, 795 heterophyiasis, 796 clinical manifestation of, 797 Phagicola spp. metacercariae in heart, 796 Pygidiopsis spp. metacercariae in kidney, 796 reagents and equipment, 801 sequences from GenBank, 800 Stellantchasmus spp. infection, 797 Human adenovirus illnesses associated with, 24 serotype classification, 24

I Immuno PCR (IPCR), 9 inlA primers for species-specific confirmation of L. monocytogenes, 216 Invasins role in aeromonad virulence, 278 Isospora spp. clinical manifestations and, 719–720 conventional laboratory diagnosis of duodenal content, 721 histological sample, 721 stool samples, collection and preservation of, 720–721 detection procedure, 727

Index diagnostic techniques histopathology, 723 stool examination, 721–723 extraintestinal tissue cysts of, 719 gametogony, 719 I. belli conventional diagnosis of, 718 description of characteristic oocysts of, 717, 719 extraintestinal tissue cysts of, 719 infection with, 719–720 life cycle of, 718 life cycle of, 718–719 molecular diagnosis of, 724 nested PCR amplification, agarose gel electrophoresis of, 727 Oocysts of, 719 RNA cistron, representation of, 727 sample preparation DNA extraction method, 726–727 stool samples, disruption of, 725–726 sporogony, 718 sporozoites, 718–719 transmission of, 719

K Klebsiella spp., 391–392 classification, 392 detection techniques PCR amplification, 398–399 rpoB gene fragments, sequence analysis of, 399 as enteropathogen, 394 as foodborne pathogen, 394 habitat and epidemiology of, 392 healthcare associated infections caused by, 393–394 isolation and biochemical identification, 394 K. pneumoniae community acquired pneumonia (CAP), 392–393 infections, 392 molecular identification, 394–395 molecular typing, 395–396 PCR and sequencing methods, 396 PCR DNA fragments, gel electrophoresis of, 397 phylogenetic analysis, 400 and Raoultella, classification of, 392 rhinoscleroma, 393 sample collection and preparation DNA isolation and, 397–398 sample processing and culture, 397 Kocuria spp. catheter-related bacteremia, 202 classification of, 201 clinical relevance, 201 diagnostic methods phenotypic methods, 202 drug susceptibilities of, 202 and food products, 202 identification and susceptibility test systems, 202–203 Kocuria kristinae acute cholecystitis and, 202 phenotypic variability, 202 for standardize quality of fermented sausages, 202 Micrococcaceae family, 201 misidentification of, 203 PCR amplification of 16S RDNA region, 204 16S ± 23S RDNA intergenic region, 204 ribotyping, 203–204 Kuru transmissible agents in dementias, 113

873

Index L β-Lactams in gram-negative bacilli, 462 production by Serratia spp., 463 S. marcescens, resistance mechanisms of β-lactamses, 461–462 gram-negative bacteria, outer membrane of, 462 S-Layer proteins role in aeromonad virulence, 278 Lipase-encoding genes role in aeromonad virulence, 278 Lipopolysaccharide role in aeromonad virulence, 278 Listeria spp. classification of, 207 detection techniques identities and sequences of internalin gene primers for species-and virulence determination, 216 identities and sequences of primers for serogrouping, 216 identities and sequences of primers for species-specific identification, 215 multipex PCR determination of virulence, 215 multipex PCR identification, 214 multiplex PCR differentiation of serogroups, 215 diagnosis agglutination and enzyme-based immunoassay (EIA), 212–213 biochemical tests, 212 molecular techniques, 213 in vitro culture isolation, 212 in vivo assays, 213 distribution in environment, 208 fish and seafoods, isolation from, 209 genes for virulence-associated proteins, 210 genomes comparison, 211–212 growth of, 208 infection abortion/septicaemia, 209 early-onset fetomaternal/neonatal listeriosis, 210 food related human listeriosis, outbreaks of, 209 granulomatosis infantiseptica, 210 mode of, 210 processed and ready-to-eat (RTE), 209 Listeria primers for species specific identification, 212 Listeria monocytogenes serovars human listeriosis, 208 intracellular mobility and cell-to-cell spread, 210 and relative virulence, 209 Listeria somatic (O)/flagellar (H) antigens and antisera compositions of, 208 serological techniques for interaction between, 207 sample preparation bacterial isolation, 213–214 DNA extraction, 214 whole genome sequences of, 210 Longicatenatae, 521 Loop mediated amplification (LAMP), 7 Lymphoid cell depletion and astroviruses infection, 36

M Map animal pathogen and Crohn’s disease in humans, 230 detection of bacteriophage-based rapid methods, 232 internal amplification control (IAC), 235 microarray technology, 235 NASBA, 235 nested/semi-nested PCR, 233 phenotypic techniques, 231–232 primers for targeting, 234–235 probs for, 236 real-time PCR methods, 233

single PCR, 232–233 detection techniques DNA extraction protocol, 238–239 IS900 single PCR procedure, 239 in food, 231 genome sequence, 229 Johne’s disease in ruminants, 230 reagents and equipment, 237 sample preparation, 237–238 in water, 231 Metagonimus spp. detection procedures Karyotyping of, 810 PCR-RFLP, 808–809 RAPD analysis, 809 28S D1 rDNA and mtCOI PCR, sequencing of, 809–810 simple sequence repeat-PCR, 809 Heterophyidae family, 805 identification and diagnosis, 807 medical importance, 807 metagonimiasis, 805 M. miyatai biology of, 806 classification and morphology of, 806 M. takahashii biology of, 806 classification and morphology of, 806 M. yokogawai biology of, 806 classification and morphology of, 806 pathogenicity, 806–807 reagents and equipment, 808 sample collection and preparation DNA extraction, 808 Micrococcus spp. aroma and flavor, development of, 222 biogenic amines (BA) production, 222 molecular detection, 225 poisoning, 223 diagnosis, 223 food matrices preparation, 223 dairy products, 224 fish and sea food, 224 meat and meat products, 224 food spoilage by, 222 habitats of, 222 identification of, 221 M. luteus biovar II strains, 225–226 characteristics differentiating Kocuria varians and, 225 house-keeping genes, 225–226 PCR detection, 224–225 variability in physiological traits, 221 M. lylae isolated from human skin habitat, 221 phenotypic differentiation between, 222 quinone system and peptidoglycan type, 221 Microsporidia organisms, 691 detection techniques multiplex nested PCR protocol, 696 oligonucleotide primers and detection probes, 697–698 real-time PCR method, 696–697 diagnosis microscopy by, 693 serological methods, 694 in vitro culture, 693–694 in vivo assays, 694 E. bieneusi genotypes, 693, 697 E. cuniculi strain I geographical distribution, 693

874 infection in humans and animals, 692 life cycle, 692 microsporidial spores, 692 molecular techniques, 694 genotyping, 695 species-specific detection, 694 susceptibility testing, 695 sample preparation DNA isolation, 695 Micro total analysis systems (µTAS), 9 Minitek system (BBL), 568 Molecular beacons, 6–7 Molecular Beacon technologies, 5 Multiplex PCR for Listeria monocytogenes speciation and virulence determination, 7 Multiplex PCR method, use of genes in, 537–538 Mycobacterium avium subsp. paratuberculosis, IS900 as insertion element, 13 Mycobacterium spp. growth rate, 229 identification of, 229 Map-specific PCR targets, 229–230 Mycobacteriaceae family, 229

N NADH dehydrogenase genes from mt DNA as target for detection of helminths, 13 Noncatenatae, 521 Noroviruses (NoVs) classification of, 75 cloning and sequencing of genome, 77 detection in food amplification and detection, 80 amplification and inhibition controls, 85 clarification of water-phase by extracting with chloroform-butanol mixture, 80 elution and concentration, 77 EU legislation, 82 flow chart of, 84 GI and GII assay, 81 mastermix and RT-PCR conditions, 85 quality assurance (QA)/quality control (QC), 82 real-time RT-PCR primers and probes targeting ORF 1-ORF 2 junction, 81 RNA extraction, 80 RT-PCR amplicons, 80 sample processing, 77 in shellfish, 78–79 in soft fruits, 84–85 TaqMan real-time protocol, 84 in vegetables and berries, 78–79 diagnosis, 77 foodborne outbreaks in U.S., 75 genome with open reading frames (ORFs), 76 ORF1–ORF2 junction region, 80 incubation period, 76 infection 1,2-fucosyltransferase FUT2 gene, susceptibility of, 77 human histoblood group antigens (HBGAs) and, 76–77 source of, 76 ready-to-eat (RTE) foods, 75 sample preparation and RNA extraction, 82 reagents and equipments, 83 Nucleic acid based, pathogen detection techniques fluorescence in situ hybridization (FISH), 8 immunological detection methods, 8–9 isothermal amplification, 7–8 microarray detection, 8 polymerase chain reaction (PCR)

Index double/single stranded (ss) DNA sequences in vitro, amplification of, 3 multiplex PCR assays, 7 practical considerations for, 4 real-time PCR systems, 6–7 representation and detection protocols, 5 reverse transcriptase PCR, 7 steps used for, 5 Nucleic acid sequence based amplification (NASBA), 7 for detection Hepatitis A virus (HAV), 8 Vibrio cholerae, 8

O OIE Manual for Diagnostic Tests and Vaccines for Terrestrial Animals, 55 Oncosphere-specific protein tso31 as detection target for helminth Taenia, 14 Opisthorchis spp. biology and life cycle, 814–815 cholangiocarcinoma (CCA), 813 liver fluke induced, 817–818 nitrative and oxidative DNA damage, 818 classification, 813–814 clinical manifestation and morbidity, 816–817 diagnosis immunological techniques, 820–821 molecular techniques, 821 parasitological techniques, 819–820 epidemiology, 819 geographical distribution, 814 host immune response, 816 opisthorchiasis, 820 praziquantel chemotherapy, 819 Opisthorchiidae family, 813 O. viverrini antigens of, 821 commercial kit, DNA preparation, 821–822 CTA B method, DNA preparation, 822 endogenous nitrosation, 818 host, 816 infection, 815 8-oxodG levels and, 818 stool samples PCR detection by pOV-6 primers, 822–823 stool samples PCR detection by trem25-F and OV25-4R primers, 823 pathogenesis and pathology, 815–816 prevention, and control, 819 Orthomyxoviruses, as biological agents of biohazard class 2, 56 Outer membrane proteins (OMP) role in aeromonad virulence, 278

P Paragonimus spp. biology and pathogenesis, 829–830 classification and morphology P. africanus, 829 P. kellicotti, 829 P. mexicanus, 829 P. skrjabini and P. miyazakii, 829 P. tuanshanensis, 829 P. uterobilateralis, 829 P. westermani, 828–829 detection procedures, 833 cycle sequencing, 834 ethanol precipitation of cycle sequencing product, 834–835 PCR amplification, 834 purification of PCR fragment from gel, 834 diagnosing infection enzyme linked immunosorbent assay (ELISA), 832

875

Index immunoblot assays, 832 immunological techniques, 830–832 intradermal test, 831–832 DNA sequence data and phylogenetic analysis, 828 paragonimiasis/endemic hemoptysis, 827 diagnostic methods, 830–831 sample collection, 832–833 DNA extraction from eggs, 833 Penicillium spp., 593 classification of, 594 detection procedures PCR detection of Penicillium species, 599 human diseases and, 595–596 laboratory diagnosis conventional techniques, 596 molecular techniques, 596–598 morphology of, 594 mycotoxinogenic foodborne Penicillia human health and, 595 OTA, 595 patulin, 595 nature and habitat, 594 PCR detection, 599 OTA-producing Penicillium, 599–600 patulin-producing Penicillium, 600 samples preparation culture preparation for toxin detection, 598 DNA extraction from fungal Cultures, 598–599 isolation from air, 598 isolation from food samples, 598 Pharyngoconjunctival fever (PCF), 25; see also Adenoviruses Phenotypic switching”, 552 Plate count technique, 534 Plesiomonas spp. equipment and supplies, 411–412 habitat, 406 infection control, 409 pathogenicity and virulence factors, 408 PCR assay and visualization, 412 P. shigelloides conventional diagnosis, 409–410 extraintestinal infections, 407 in foods, factors affecting, 406–407 gastroenteritis, 407 hemolytic activity, 409 hugA gene, amplification of, 412 invasiveness, 408–409 molecular diagnosis of, 410–411 PCR assays developed for, 411 plasmids, 409 tetrodotoxin (TTX), 409 toxins, 408 typing, 411 reagents, 411 sample collection, 412 taxonomy of, 405–406 Polymerase chain reaction (PCR) for amplification of double/single stranded (ss) DNA sequences in vitro, 3 amplification efficiency, 4 annealing time, 4 hairpin primers, 7 microchip PCR systems, 9 PCR-RFLP (restriction fragment length polymorphism), 9 thermostable polymerase, use of, 4 in food industry, 555 Prion diseases, see Transmissible spongiform encephalopathies (TSEs) Proteases role in aeromonad virulence, 278 Pseudomonas spp. detection techniques, 440

P. aeruginosa acquisition of antibiotic resistance in, 432 detection in food products, 436–438 environmental sources of, 431–432 hospitalized patients, risk of acquisition of, 432–436 in human activity environments, selective advantage for, 432 identification in hospitals clinical laboratories, 439 novel nonantibiotic strategies to, 436 virulence potential, assessment of, 439–440 sample preparation bacterial DNA isolation, 440 Pulsed field gel electrophoresis (PFGE), 9

R Real-time PCR (rt-PCR), for fungal detection, 536 Real-time PCR systems nonspecific detection, 6 sequence specific detection, 6 Reverse transcriptase PCR, 7 Rhodotorula spp., 603 affinity for plastic, 605 conventional identification, of yeasts, 606–607 detection methods and procedures DNA extraction, 609 identification and genotyping, by MSP-PCR typing method, 609–610 rDNA sequencing analysis, 611–612 by RFLP-ITS method, 611 sample collection and preparation, 609 by species-specific PCR primers, 610–611 environmental distribution of, 604–605 future research areas, 613 medical importance, 605–606 molecular diagnostic methods, 607–609 occurrence in food and beverages, 605 role in human infections, 605 taxonomy of, 603–604 treatment of infection, 606 type strains of medically relevant Rhodotorula species, 612 Rolling circle amplification (RCA), 7 Rotaviruses detection techniques primers and probes, 97 real-time RT-PCR detection of groups A and C, 96–97 RT-PCR detection of G and P genotypes, 96 VP4 and VP7 primers for, 96 diagnosis acid adsorption–alkaline elution and reconcentration, 95 antigen detection techniques, 94 enzyme immunoassay, 94 molecular procedures, 95 PCR assays, 95 phenotypic techniques, 94 family Reoviridae, 91 genetic changes by sequential point mutations, 93 G-type classification, 92 infection clinical symptoms, 93 gastroenteritis with diarrhea, 93 replication, 93 severity of, 94 transmitted by fecal-oral route via contact with contaminated hands, 93 infectivity of, 92 nonstructural protein (NSP), 91 overlapping reading frames, 91 RNA-binding proteins, 92

876 serogroups, 92 serotypes and genotypes, 91–92 rpsU-dnaG-rpoD region, DNA and RNA synthesis, 13

S Saccharomyces cerevisiae, 619–620 colony formation, 621 detection methods and procedure identification, by restriction of amplified 5.8S-ITS region of ribosomal rRNA operon, 632 microbial cells separation, from bakery dough, 631 reagents and equipment, 627 real-time PCR, detection by, 631–632 sample treatment, 627 typing by microsatellite amplification, 632–633 DNA extraction from human fecal samples, 627, 629–631 pure cultures, 627 from wine, 631 food industry, use in, 620 future investigation approach, 633 genetic typing methods and, 625 life cycle of, 620 molecular techniques for detection and identification, 625–628 microsatellite application, 626 occurrence in nature, 620 as pathogen, 622–624 retrotransposon types, 620 Saccharomyces complex, 619 S. boulardii as variant of, 621–622 virulence factor in pathogenic strains, 624 wine fermentation, 620 Salmonella spp. biology of serotyping, 447–448 detection TaqMan Real-Time PCR method for, 454–455 diagnostics of biochemical tests and serotyping, 450 culture based method, 450 modified conventional methods, use of, 450 real-time PCR method for, 449 Enterobacteriaceae family, 447 epidemiology cyclic between humans and animals, transmission of, 448 S. enteritidis and S. typhimurium, 448 S. enteritidis, reservoir for, 448–449 infection route, 448 molecular diagnostics, 450 immunological methods, 451–452 nucleic acid-based methods, 452–453 pathogenesis infections, nature and severity of, 448 type III secretion systems (TTSS) for, 448 sample collection and preparation, 453–454 Sapoviruses classification family Caliciviridae, 101 Star-of-David, animal calicivirus morphology, 101 detection techniques ELISA detection system, 107 by EM, 106–107 integrity and morphology, 107 nested PCR, 108 phylogenetic analysis, 109 QIAamp Viral RNA Protocol, 107 real-time RT-PCR method, 108–109 reverse transcription (RT), 107–108 RNA extraction, 107

Index diagnosis enzyme immunoassays (EIAs), 103 nested PCR, 103 real-time RT-PCR methods, 103 genome genogroups (GI–GV), 102 open reading frames (ORFs), 102 phylogenetic analysis, 102 RdRp-capsid junction, 102 SimPlot software, 102 infections gastroenteritis, 101 sporadic cases of acute gastroenteritis, 103 virulence prevalence, 101 molecular detection nested RT-PCR by, 104 reagents and equipments, 105 RT-PCR and identification by sequence analysis, 103–104 sample preparation influent and effluent water, 105–106 oysters, 106 primary-treated domestic wastewater, 105 river water, 106 seawater, 106 secondary-treated effluent and river water, 105 shellfish, 106 stool samples, 106 subgenomic-like construct, expression of, 103 virus-like particles (VLPs), formation of, 103 Sarcocystis spp. detection procedures, 734 18S rDNA sequencing, 735 DNA extraction sarcocysts and cell cultures from, 734 single tube extraction from oocysts, 733–734 in vitro excystation method, 734 feeding contaminated meat, 731 mature, excreted oocysts of, 731 sample collection and preparation preliminary visual screening for sporocysts, 732 sporocysts isolation, 732–733 Serratia spp. classification of, 460 laboratory identification of, 465 PCR and restriction enzyme digestion PCR amplification, 466 restriction endonuclease digestion, 466 sample preparation, 466 real-time PCR amplification and detection, 466 sample preparation, 466 S. marcescens antimicrobial resistance, occurrence of, 461 multidrug efflux pumps identification in, 464 opportunistic pathogen, 460–461 presence of, 459–460 resistance mechanisms, aminoglycosides, 464–465 resistance mechanisms, β-lactams, 461–462 resistance mechanisms, fluoroquinolones, 462–464 Shigella spp. actin assembly, 474 contaminations of, 472 detection selected rapid methods for, 477 diagnosis conventional detection in food, 475–476 molecular detection in food, 477–478 FDA investigation of foodborne pathogens, 473 foodborne outbreaks of, 472–473 FoodNet data, 473 hemolytic uremic syndrome (HUS), 475

877

Index icsA (virG) and icsA gene, 474 infection route, 474 macropinocytosis, 474 medical importance, 475 molecular evolutionary relationship between enteroinvasive E. coli (EIEC) and, 471–472 nested PCR detection of electrophoresis of, 480 primary PCR, 480 primer solutions, 479–480 outbreaks and foods, 472 pathogenesis, 473 IcsA is distribution, 474–475 intra and intercellular spreading, 474 macropinocytosis involvment, 474 Shiga-like toxin (SLT) production and, 474 Shiga toxin (STX), 475 PCR analyte and bacteriological analysis, preparation of, 478 PCR template, preparation of, 479 sample enrichment, 478 S. boydii serotype, 472 S. dysenteriae serotype, 472 serogroups of, 473 S. flexneri infections, 475 LPS mutant, 474 serotype, 472 shigellosis treatment rehydration and antimicrobial therapy, 475 suspect colonies, identification of, 478–479 Wiskott–Aldrich syndrome protein (N-WASP) and, 474 Slow virus diseases, 113 diagnostic methods enzyme-linked immunosorbent assay (ELISA) techniques, 114 histological and immunohistochemical (IHC) methods, 117 western blot method, 117 virus-induced, 114 Staphylococcus spp. coagulase-negative (CNS), 245 colonies of, 245 detection techniques, 253 primer identities, 254 reagents and equipment, 252 sample collection meat sampling, 253 milk sampling, 252–253 sample pretreatment DNA extraction, 253 S. aureus coagulase-positive (CPS), 245 endemic strains, 246 in food, 246–247 multiplex PCR, 249–250 nuc gene as thermostable nuclease, 13 real-time PCR, 250–251 reservoirs of, 245 risk assessment in foodstuffs, 247 standard PCR protocols, 247–249 Staphylococcal food poisoning (SFP) in food and food products, 246 outbreaks, 246 symptoms of, 246 virulence factor SE characteristics of, 246 types of, 246 Strand displacement amplification (SDA), 7 Streptococcus spp.

classification of, 259 Group A streptococci (GAS), 260 Group B streptococci (GBS), 260 Groups D, E, and F streptococci, 260 M-types GAS, 260 serological classification system, 260 dectection techniques composition of primers used in SA g PCR, 267 multiplex PCR for superantigen, 265–266 PCR for emm-typing GAS, 265 diagnosis GAS quick tests, 265 Lancefield agglutination test, 261 molecular techniques, 265 pneumococcus quick test, 265 foodborne infections, 262–264 outbreaks, 261 invasive pneumococcal disease (IPD) in USA, 260 Pneumococcal infections, 260 S. agalactiae neonatal infections, 261 perinatal infections, 261 puerperal sepsis, 259 sample collection, 265 scarlet fever, 261 serotype, determination of, 260 skin infection, 261 S. mitis mosaic genes and, 260 S. oralis mosaic genes and, 260 S. pneumoniae colony in human nasopharynx, 261 meningitis and pneumonia, 259 streptococcal superantigens (SAg) infections, 261 streptococcal toxic shock-like syndrome (STSS), 259 streptococcal tonsillopharyngitis, 259 streptolysins O and F, toxins, 261 Superoxide dismutase role in aeromonad virulence, 278 Surface-enhanced Raman scattering (SERS), ssDNA detection, 10 Surface plasmon resonance (SPR) sensors for measurement of refractive index, 8 Systematic evolution of ligands by exponential enrichment (SELEX), 10

T Taenia spp. biology and pathogenesis of foodborne, 840 classification, 840 cysticercosis and echinococcosis, 839 detection procedures multiplex PCR, 845–846 PCR product, sequencing of, 846–847 diagnosis, techniques for, 839–840 histopathological specimens for molecular identification, 841 human sparganosis, 839 human taeniid species molecular probes for identification of, 844–845 multiplex PCR for, 846 immunological approaches immunodeficient mice as alternative intermediate hosts, usefulness of, 842 life cycle of, 840 metacestodiases, 839 molecular and immunological techniques, relative performance of, 842–843 molecular tools for prevention of NCC, 840 loop-mediated isothermal amplification (LAMP) tool, 840–841

878 neurocysticercosis (NCC), 839 in humans, 842 racemose cysticercosis, 841 sample collection and preparation, 843, 845 taeniid proglottids expelled from humans, molecular diagnostic techniques, 841 tapeworm infections, 839 T. asiatica scoleces of, 841 T. asisatica cysticercosis in pigs, 842 T. saginata bovine cysticercosis in cattle, 842 cysticerci of, 843 distribution of, 840 human taeniasis and cysticercosis of, 840 T. solium cysticercosis in, 842 distribution, 840 genotypes of, 841 human taeniasis and cysticercosis of, 840 scoleces of, 841 zoonotic taeniid infections, 842 Targets for detection techniques genes for surface expressed markers, 13–14 insertion elements, 13 mitochondrial genes, 13 nonviral targets cytoskeletal proteins, 12 ribosomal RNA (rRNA) genes, 11–12 RNA targets, 11 unique genes and sequences, 12–13 viral, 10–11 structural genes, 11 virulence and toxin genes, 12 Toxoplasma B1 gene as detection target, 13 Toxoplasma spp. bradyzoite, 742–743 cats and mice, bioassay in, 744–745 detection procedures multilocus nested PCR-RFL P genotyping, 749–750 nPCR, 747–748 by real-time PCR, 748–749 diagnosis enzyme-linked immunoabsorbent assay (ELISA), 743–744 indirect fluorescent antibody test (IFA), 743 indirect hemagglutination test (IHA), 743 modified direct agglutination test (MAT), 744 Sabin–Feldman dye test (DT), 743 epidemiology and pathogenesis, 743 molecular techniques multilocus nested PCR-RFLP genotyping, 745–746 nested PCR (nPCR) of repetitive, DNA sequences, 745 reagents and equipment, 746 real-time PCR of repetitive DNA sequences, 745 multilocus nested PCR-RFL P markers, summary of PCR primers for, 750 multilocus PCR-RFL P genotyping results, summary of, 751 PCR-RFLP gel image, 751 sample preparation DNA from animal tissues, extraction, 746–747 oocysts from water samples and extract, purification, 747 T. gondii host genetic makeup and parasite genotypes, 745 infectious stages of, 741–742 isolation of, 744 life cycle and, 741–742 oocysts, 742–743 Transmissible spongiform encephalopathies (TSEs), 113 detection techniques, 120–121 abnormal isoform of prion protein, 122

Index diagnostic methods, 117 capillary gel-electrophoresis, 119 conformation-dependent immunoassay (CDI), 119 fluorescence correlation spectroscopy (FCS), 119 modified western blotting method for prion detection, 118 multi-spectral ultraviolet fluorescence spectroscopy (MUFS), 119 tool for, 119 visible and near-infrared (Vis-NIR) spectroscopy, 119 mutations responsible for inherited genetic forms, 114 genes, 115 sample collection and preparation, 120 Trichinella spp. amplicon sizes by PCR-RFL P, features of, 852 biology of, 853 detection procedures multiplex PCR, 857 encapsulated clade T. britovi, 852 T. murrelli, 853 T. nativa, 852 T. nelsoni, 853 Trichinella T6, 852 Trichinella T8, 852–853 Trichinella T12, 853 expected amplification products, dimension of, 860 hosts for, 852 infection, diagnosing methods for, 854–855 molecular probes, 855 multiplex-PCR, 855 PCR-restriction fragment length polymorphism, 855 random amplified polymorphic DNA (RAPD), 855 reverse line blot hybridization (RLB), 855–856 single-strand conformational polymorphism (SSCP), 855 medical importance, 854 non-encapsulated clade T. papuae, 853 T. pseudospiralis, 853 T. zimbabwensis, 853 pathogenesis, 853 infection pathological picture of, 854 larvae transformation, 854 PCR amplification of single larvae, photograph of, 859 sample collection and preparation DNA purification from single larvae, 857 hosts and preferential muscles, 856 parasite isolation and preservation, 856–857 SSCP approach for, 860 T. britovi and Trichinella T8, PCR amplification of single larvae of, 860 and Trichinella T9, PCR-RFLP to distinguish between, 858–859 and Trichinella T8, PCR to distinguish between, 859–860 zoonosis trichinellosis, 851 Trichothecenes, 579 β-Tubulin as targets, 12; see also Targets for detection techniques Turkey astrovirus 2 (TAstV-2) infection, mild crypt hyperplasia, 36 Turkey (TAstVs) serotypes of astroviruses, 35

U U.S. foodborne illness outbreaks, statistics, 2

V Vibrio spp. biology and pathogenesis, 486 halophilic vibrios, 488 classification of, 486 detection techniques PCR assays, 493 primers and probes, 494

879

Index diagnosis, 488 DNA-DNA colony hybridization in, 489–490 PCR technology, applications of, 490 real-time PCR using, 491 reverse transcriptase PCR (RT-PCR), application of, 491–492 SYBR Green I-based, 491 TaqMan quantitative PCR (qPCR), study, 490–491 V. vulnificus in oysters, 491 outbreaks, 485 transmission, source, 485 V. cholerae biology and pathogenesis, 487 real-time PCR assays, 493, 495 sample preparation, 492 serotypes, 487 V. parahaemolyticus, 489–490 clinical manifestations of, 487 foodborne gastroenteritis, 487 H antigen, 487 Kanagawa phenomenon, 487 outbreaks, 487 pathogenic and nonpathogenic strains, 487 real-time PCR assays, 495 sample preparation, 493 serotype, 487 wound infection and septicemia, 487 V. parahaemomlyticus biology and pathogenesis, 487 V. vulnificus biology and pathogenesis, 488 liver disease and, 488 prevalence of, 488 real-time PCR assays, 493, 495 sample preparation, 492–493 Voges-Proskauer reaction, biochemical identification of Aeromonas spp., 279

W Waldenstrom’s macroglobulinaemia, 37

Y Yersinia spp., 502 detection methods

amplification facilitators, use of, 510 enrichment cultivation step, 510 equipment and materials, 508 procedure, 510 sample preparation, 509–510 YPCE, use of, 510 detection procedure, 510 confirmation of viable target bacteria, 511 DNA extraction, 511 enrichment of food samples, 510–511 pipetting scheme for detection of ail-gene, 512 primers and probes used, 512 quality assurance and control measure, 511 real-time PCR, 511–512 Y. enterocolitica, 509–510 Y. pestis, 504, 510 Y. pseudotuberculosis, 506, 510 genome comparison of Yersinia strains, 502 Y. enterocolitica biotypes, 506 characteristics of, 505 detection by culture method and PCR, 507–508 incidences of, 506 isolation of serotypes, 507 molecular detection methods for, 509 pigs, as carriers of pathogenic strains, 507 serotypes, 505 symptoms of infection, 506 transmission of, 507 virulence factors, 506 Y. pestis disease caused by, 502–503 epidemiology, 503 isolation and molecular detection methods of, 503–504 life-cycle, 503 plasmids, 503 pneumonic plague, transmission of, 503 Y. pseudotuberculosis, 503 epidemiology, 505 incidence and prevalence of, 505 isolation rates from animal sources, 505 O-serotypes, 504–505 PCR-based detection methods for, 505–506 spread of infection, 505 symptoms of infection, 503 virulence factors, 503–504