Molecular Detection of Human Bacterial Pathogens

EDITED BY

DONGYOU LIU

MOLECULAR DETECTION OF HUMAN BACTERIAL PATHOGENS

MOLECULAR DETECTION OF HUMAN BACTERIAL 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 © 2011 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Version Date: 20110629 International Standard Book Number-13: 978-1-4398-1239-6 (eBook - PDF) 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. Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

This volume is dedicated to a group of bacteriologists whose insight, knowledge, and expertise have made the all-inclusive coverage of major human bacterial pathogens a reality.

Contents Preface���������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������� xv Editor����������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������xvii Contributors�������������������������������������������������������������������������������������������������������������������������������������������������������������������������������xix Chapter 1 Introductory Remarks��������������������������������������������������������������������������������������������������������������������������������������������� 1 Dongyou Liu

Section I  Actinobacteria Chapter 2 Actinomadura�������������������������������������������������������������������������������������������������������������������������������������������������������� 13 Martha E. Trujillo Chapter 3 Actinomyces����������������������������������������������������������������������������������������������������������������������������������������������������������� 23 Debarati Paul, D. Madhusudan Reddy, Dipalok Mukherjee, Biswajit Paul, and Debosmita Paul Chapter 4 Atopobium������������������������������������������������������������������������������������������������������������������������������������������������������������� 31 Piet Cools, Hans Verstraelen, Mario Vaneechoutte, and Rita Verhelst Chapter 5 Bifidobacterium����������������������������������������������������������������������������������������������������������������������������������������������������� 45 Abelardo Margolles, Patricia Ruas-Madiedo, Clara G. de los Reyes-Gavilán, Borja Sánchez, and Miguel Gueimonde Chapter 6 Corynebacterium�������������������������������������������������������������������������������������������������������������������������������������������������� 59 Luis M. Mateos, Michal Letek, Almudena F. Villadangos, María Fiuza, Efrén Ordoñez, and José A. Gil Chapter 7 Cryptobacterium��������������������������������������������������������������������������������������������������������������������������������������������������� 75 Futoshi Nakazawa Chapter 8 Gardnerella����������������������������������������������������������������������������������������������������������������������������������������������������������� 81 Rita Verhelst, Hans Verstraelen, Piet Cools, Guido Lopes dos Santos Santiago, Marleen Temmerman, and Mario Vaneechoutte Chapter 9 Gordonia��������������������������������������������������������������������������������������������������������������������������������������������������������������� 95 Brent A. Lasker, Benjamin D. Moser, and June Brown Chapter 10 Micrococcus and Kocuria������������������������������������������������������������������������������������������������������������������������������������111 Dongyou Liu Chapter 11 Mycobacterium����������������������������������������������������������������������������������������������������������������������������������������������������� 117 Shubhada Shenai and Camilla Rodrigues vii

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Chapter 12 Nocardia�������������������������������������������������������������������������������������������������������������������������������������������������������������� 127 Veronica Rodriguez-Nava, Frédéric Laurent, and Patrick Boiron Chapter 13 Propionibacterium���������������������������������������������������������������������������������������������������������������������������������������������� 137 Andrew McDowell and Sheila Patrick Chapter 14 Rhodococcus������������������������������������������������������������������������������������������������������������������������������������������������������� 155 Debarati Paul, Dipaloke Mukherjee, Soma Mukherjee, and Debosmita Paul Chapter 15 Saccharopolyspora���������������������������������������������������������������������������������������������������������������������������������������������� 169 Caroline Duchaine and Yvon Cormier Chapter 16 Streptomyces��������������������������������������������������������������������������������������������������������������������������������������������������������175 Angel Manteca, Ana Isabel Pelaez, Maria del Mar Garcia-Suarez, and Francisco J. Mendez Chapter 17 Tropheryma����������������������������������������������������������������������������������������������������������������������������������������������������������181 Shoo Peng Siah, Han Shiong Siah, and Dongyou Liu Chapter 18 Tsukamurella������������������������������������������������������������������������������������������������������������������������������������������������������� 189 Mireille M. Kattar

Section II   Firmicutes and Tenericutes Bacilli Chapter 19 Abiotrophia��������������������������������������������������������������������������������������������������������������������������������������������������������� 201 Marco Arosio and Annibale Raglio Chapter 20 Aerococcus�����������������������������������������������������������������������������������������������������������������������������������������������������������211 Debarati Paul, D. Madhusudan Reddy, and Dipalok Mukherjee Chapter 21 Bacillus�����������������������������������������������������������������������������������������������������������������������������������������������������������������219 Noura Raddadi, Ameur Cherif, and Daniele Daffonchio Chapter 22 Enterococcus������������������������������������������������������������������������������������������������������������������������������������������������������� 231 Teresa Semedo-Lemsaddek, Paula Lopes Alves, Rogério Tenreiro, and Maria Teresa Barreto Crespo Chapter 23 Granulicatella����������������������������������������������������������������������������������������������������������������������������������������������������� 249 Sheng Kai Tung and Tsung Chain Chang Chapter 24 Lactobacillus������������������������������������������������������������������������������������������������������������������������������������������������������� 257 Ester Sánchez and Yolanda Sanz

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Chapter 25 Leuconostoc�������������������������������������������������������������������������������������������������������������������������������������������������������� 269 María Mar Tomás and Germán Bou Chapter 26 Listeria���������������������������������������������������������������������������������������������������������������������������������������������������������������� 279 Dongyou Liu and Ting Zhang Chapter 27 Paenibacillus������������������������������������������������������������������������������������������������������������������������������������������������������� 295 Matthew R. Pincus, Nicholas Cassai, Jie Ouyang, and Sharvari Dalal Chapter 28 Staphylococcus���������������������������������������������������������������������������������������������������������������������������������������������������� 307 Paolo Moroni, Giuliano Pisoni, Paola Cremonesi, and Bianca Castiglioni Chapter 29 Streptococcus������������������������������������������������������������������������������������������������������������������������������������������������������ 323 Judith Jansen and Lothar Rink

Clostridia Chapter 30 Acidaminococcus������������������������������������������������������������������������������������������������������������������������������������������������ 339 Hélène Marchandin and Estelle Jumas-Bilak Chapter 31 Anaerococcus, Parvimonas, and Peptoniphilus������������������������������������������������������������������������������������������������� 349 Dongyou Liu and Frank W. Austin Chapter 32 Catabacter����������������������������������������������������������������������������������������������������������������������������������������������������������� 361 Susanna K.P. Lau and Patrick C.Y. Woo Chapter 33 Clostridium��������������������������������������������������������������������������������������������������������������������������������������������������������� 367 Juan J. Córdoba, Emilio Aranda, María G. Córdoba, María J. Benito, Dongyou Liu, and Mar Rodríguez Chapter 34 Dialister��������������������������������������������������������������������������������������������������������������������������������������������������������������� 381 Ana Paula Vieira Colombo, Renata Martins do Souto, and Andréa Vieira Colombo Chapter 35 Eubacterium�������������������������������������������������������������������������������������������������������������������������������������������������������� 391 Johanna Maukonen and Maria Saarela Chapter 36 Finegoldia����������������������������������������������������������������������������������������������������������������������������������������������������������� 405 A.C.M. Veloo Chapter 37 Mogibacterium�����������������������������������������������������������������������������������������������������������������������������������������������������415 Reginaldo Bruno Gonçalves, Daniel Saito, and Renato Corrêa Viana Casarin Chapter 38 Peptostreptococcus��������������������������������������������������������������������������������������������������������������������������������������������� 423 Eija Könönen and Jari Jalava

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Chapter 39 Tannerella����������������������������������������������������������������������������������������������������������������������������������������������������������� 437 Derren Ready and Lena Ciric Chapter 40 Veillonella����������������������������������������������������������������������������������������������������������������������������������������������������������� 447 Dongyou Liu

Mollicutes Chapter 41 Mycoplasma�������������������������������������������������������������������������������������������������������������������������������������������������������� 455 Rama Chaudhry, Bishwanath Kumar Chourasia, and Anupam Das Chapter 42 Ureaplasma��������������������������������������������������������������������������������������������������������������������������������������������������������� 469 Ken B. Waites, Li Xiao, Vanya Paralanov, and John I. Glass

Section III  Bacteroidetes, Chlamydiae, and Fusobacteria Chapter 43 Bacteroides���������������������������������������������������������������������������������������������������������������������������������������������������������� 491 Rama Chaudhry and Nidhi Sharma Chapter 44 Capnocytophaga������������������������������������������������������������������������������������������������������������������������������������������������� 501 Kazuyuki Ishihara, Satoru Inagaki, and Atsushi Saito Chapter 45 Chlamydia������������������������������������������������������������������������������������������������������������������������������������������������������������511 Jens Kjølseth Møller Chapter 46 Chlamydophila���������������������������������������������������������������������������������������������������������������������������������������������������� 523 Chengming Wang, Bernhard Kaltenboeck, and Konrad Sachse Chapter 47 Elizabethkingia, Chryseobacterium, and Bergeyella����������������������������������������������������������������������������������������� 537 Dongyou Liu Chapter 48 Fusobacterium���������������������������������������������������������������������������������������������������������������������������������������������������� 543 Dongyou Liu and Xiaoming Dong Chapter 49 Leptotrichia and Leptotrichia-Like Organisms�������������������������������������������������������������������������������������������������� 555 Emenike Ribs K. Eribe and Ingar Olsen Chapter 50 Porphyromonas�������������������������������������������������������������������������������������������������������������������������������������������������� 567 Stefan Rupf, Wolfgang Pfister, and Klaus Eschrich Chapter 51 Prevotella������������������������������������������������������������������������������������������������������������������������������������������������������������ 585 Mario J. Avila-Campos, Maria R.L. Simionato, and Elerson Gaetti-Jardim Jr.

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Section IV  Proteobacteria Alphaproteobacteria Chapter 52 Anaplasma����������������������������������������������������������������������������������������������������������������������������������������������������������� 601 Marina E. Eremeeva and Gregory A. Dasch Chapter 53 Bartonella������������������������������������������������������������������������������������������������������������������������������������������������������������� 617 Alessandra Ciervo, Marco Cassone, Fabiola Mancini, and Lorenzo Ciceroni Chapter 54 Brucella��������������������������������������������������������������������������������������������������������������������������������������������������������������� 629 Sascha Al Dahouk, Heinrich Neubauer, and Herbert Tomaso Chapter 55 Ehrlichia�������������������������������������������������������������������������������������������������������������������������������������������������������������� 647 Jere W. McBride, Juan P. Olano, and Nahed Ismail Chapter 56 Ochrobactrum����������������������������������������������������������������������������������������������������������������������������������������������������� 659 Corinne Teyssier and Estelle Jumas-Bilak Chapter 57 Orientia��������������������������������������������������������������������������������������������������������������������������������������������������������������� 671 Harsh Vardhan Batra and Diprabhanu Bakshi Chapter 58 Rickettsia������������������������������������������������������������������������������������������������������������������������������������������������������������� 683 Marina E. Eremeeva and Gregory A. Dasch

Betaproteobacteria Chapter 59 Achromobacter���������������������������������������������������������������������������������������������������������������������������������������������������� 703 Beth Mutai and Yi-Wei Tang Chapter 60 Bordetella������������������������������������������������������������������������������������������������������������������������������������������������������������ 709 Diego Omar Serra, Alejandra Bosch, and Osvaldo Miguel Yantorno Chapter 61 Burkholderia������������������������������������������������������������������������������������������������������������������������������������������������������� 723 Karlene H. Lynch and Jonathan J. Dennis Chapter 62 Eikenella�������������������������������������������������������������������������������������������������������������������������������������������������������������� 737 Javier Enrique Botero, Adriana Jaramillo, and Adolfo Contreras Chapter 63 Kingella��������������������������������������������������������������������������������������������������������������������������������������������������������������� 745 Dongyou Liu

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Chapter 64 Laribacter������������������������������������������������������������������������������������������������������������������������������������������������������������751 Patrick C.Y. Woo and Susanna K.P. Lau Chapter 65 Neisseria�������������������������������������������������������������������������������������������������������������������������������������������������������������� 757 Paola Stefanelli Chapter 66 Ralstonia������������������������������������������������������������������������������������������������������������������������������������������������������������� 769 Michael P. Ryan, J. Tony Pembroke, and Catherine C. Adley

Gammaproteobacteria Chapter 67 Acinetobacter������������������������������������������������������������������������������������������������������������������������������������������������������ 781 Dongyou Liu Chapter 68 Aeromonas����������������������������������������������������������������������������������������������������������������������������������������������������������� 789 Germán Naharro, Pedro Rubio, and José Maria Luengo Chapter 69 Aggregatibacter��������������������������������������������������������������������������������������������������������������������������������������������������� 801 Dongyou Liu Chapter 70 Cardiobacterium��������������������������������������������������������������������������������������������������������������������������������������������������811 Efthimia Petinaki and George N. Dalekos Chapter 71 Cedecea����������������������������������������������������������������������������������������������������������������������������������������������������������������817 Maria Dalamaga and Georgia Vrioni Chapter 72 Citrobacter���������������������������������������������������������������������������������������������������������������������������������������������������������� 827 Ignasi Roca and Jordi Vila Chapter 73 Coxiella��������������������������������������������������������������������������������������������������������������������������������������������������������������� 837 Claire Pelletier Chapter 74 Enterobacter�������������������������������������������������������������������������������������������������������������������������������������������������������� 853 Angelika Lehner, Roger Stephan, Seamus Fanning, and Carol Iversen Chapter 75 Escherichia���������������������������������������������������������������������������������������������������������������������������������������������������������� 869 P. Elizaquível, G. Sánchez, and R. Aznar Chapter 76 Francisella tularensis����������������������������������������������������������������������������������������������������������������������������������������� 881 Kiersten J. Kugeler and Jeannine M. Petersen Chapter 77 Haemophilus������������������������������������������������������������������������������������������������������������������������������������������������������� 893 Dongyou Liu, Jianshun Chen, and Weihuan Fang

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Chapter 78 Klebsiella������������������������������������������������������������������������������������������������������������������������������������������������������������� 905 Beatriz Meurer Moreira, Ana Cristina Gales, Maria Silvana Alves, and Rubens Clayton da Silva Dias Chapter 79 Legionella�������������������������������������������������������������������������������������������������������������������������������������������������������������919 Dianna J. Bopp, Kimberlee A. Musser, and Elizabeth J. Nazarian Chapter 80 Moraxella������������������������������������������������������������������������������������������������������������������������������������������������������������ 929 Suzanne J.C. Verhaegh and John P. Hays Chapter 81 Pasteurella���������������������������������������������������������������������������������������������������������������������������������������������������������� 945 Francis Dziva and Henrik Christensen Chapter 82 Photobacterium��������������������������������������������������������������������������������������������������������������������������������������������������� 959 Carlos R. Osorio and Manuel L. Lemos Chapter 83 Plesiomonas�������������������������������������������������������������������������������������������������������������������������������������������������������� 969 Jesús A. Santos, Andrés Otero, and María-Luisa García-López Chapter 84 Proteus���������������������������������������������������������������������������������������������������������������������������������������������������������������� 981 Antoni Róz˙alski and Paweł Sta˛czek Chapter 85 Providencia��������������������������������������������������������������������������������������������������������������������������������������������������������� 997 Brunella Posteraro, Maurizio Sanguinetti, and Patrizia Posteraro Chapter 86 Pseudomonas���������������������������������������������������������������������������������������������������������������������������������������������������� 1009 Timothy J. Kidd, David M. Whiley, Scott C. Bell, and Keith Grimwood Chapter 87 Salmonella��������������������������������������������������������������������������������������������������������������������������������������������������������� 1023 Mathilde H. Josefsen, Charlotta Löfström, Katharina E.P. Olsen, Kåre Mølbak, and Jeffrey Hoorfar Chapter 88 Serratia�������������������������������������������������������������������������������������������������������������������������������������������������������������� 1037 Kevin B. Laupland and Deirdre L. Church Chapter 89 Shigella�������������������������������������������������������������������������������������������������������������������������������������������������������������� 1049 K.R. Schneider, L.K. Strawn, K.A. Lampel, and B.R. Warren Chapter 90 Stenotrophomonas�������������������������������������������������������������������������������������������������������������������������������������������� 1063 Martina Adamek and Stephan Bathe Chapter 91 Vibrio����������������������������������������������������������������������������������������������������������������������������������������������������������������� 1073 Asim Bej Chapter 92 Yersinia�������������������������������������������������������������������������������������������������������������������������������������������������������������� 1089 Mikael Skurnik, Peter Rådström, Rickard Knutsson, Bo Segerman, Saija Hallanvuo, Susanne Thisted Lambertz, Hannu Korkeala, and Maria Fredriksson-Ahomaa

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Contents

Epsilonproteobacteria Chapter 93 Arcobacter���������������������������������������������������������������������������������������������������������������������������������������������������������� 1111 Kurt Houf Chapter 94 Campylobacter���������������������������������������������������������������������������������������������������������������������������������������������������1125 Rodrigo Alonso, Cecilia Girbau, and Aurora Fernández Astorga Chapter 95 Helicobacter�������������������������������������������������������������������������������������������������������������������������������������������������������1141 Athanasios Makristathis and Alexander M. Hirschl

Section V  Spirochaetes Chapter 96 Borrelia��������������������������������������������������������������������������������������������������������������������������������������������������������������1155 Nataliia Rudenko, Maryna Golovchenko, James H. Oliver Jr., and Libor Grubhoffer Chapter 97 Leptospira����������������������������������������������������������������������������������������������������������������������������������������������������������1169 Rudy A. Hartskeerl Chapter 98 Treponema���������������������������������������������������������������������������������������������������������������������������������������������������������� 1189 Rita Castro and Filomena Martins Pereira

Section VI  Pan-Bacterial Detection Chapter 99 Pan-Bacterial Detection of Sepsis-Causative Pathogens���������������������������������������������������������������������������������� 1203 Roland P.H. Schmitz and Marc Lehmann Chapter 100 Metagenomic Approaches for Bacterial Detection and Identification���������������������������������������������������������������1215 Chaysavanh Manichanh

Preface Bacteria are small, unicellular organisms that are invisible to the naked eye but are nonetheless present ubiquitously and abundantly in all environments. Although a majority of bacteria are free-living and symbiotic, some are capable of leading a parasitic life, inducing a range of disease syndromes in human and animal hosts during the process. Among the most devastating bacterial pathogens, Yersinia pestis, the causative agent of bubonic plague, was responsible for three major human pandemics in history, killing 200 million people prior to the advent of antibiotics. The current, most common fatal bacterial diseases are tuberculosis (caused by Mycobacterium tuberculosis), killing about 2 million people a year alone, and cholera (caused by Vibrio cholerae). Other globally important bacterial diseases include pneumonia (caused by Streptococcus and Pseudomonas), tetanus, typhoid fever, diphtheria, syphilis, and leprosy. Traditionally, bacteria have been identified and diagnosed with the help of various phenotypic procedures, such as Gram stain, morphological, biochemical, and serological examination. Since the phenotypic techniques are often slow and lack desired specificity and reproducibility, nucleic acid amplification technologies such as polymerase chain reaction (PCR) have played an increasingly prominent role in the laboratory diagnosis of bacterial infections. Given their ability to specifically detect a single copy of bacterial nucleic acid template in a matter of hours, PCR-based assays offer unsurpassed sensitivity, specificity, accuracy, precision, and result availability for bacterial identification. The recent advances in instrumentation automation and probe chemistries have facilitated the development of real-time PCR that provides a convenient platform for high throughput detection and quantitation of bacterial pathogens in clinical specimens. Considering that numerous original molecular protocols and subsequent modifications have been described and

scattered in various journals and monographs, it has become difficult if not impossible for someone who has not been directly involved in the development of original or modified protocols to know which are most appropriate to adopt for accurate identification of bacterial pathogens of interest. The purpose of this volume is to address this issue, with international scientists in respective bacterial pathogen research and diagnosis providing expert summaries on current diagnostic approaches for major human bacterial pathogens. Each chapter consists of a brief review of the classification, epidemiology, clinical features, and diagnosis of an important pathogenic bacterial genus; an outline of clinical sample collection and preparation procedures; a selection of representative stepwise molecular protocols; and a discussion on further research requirements relating to improved diagnosis. This book represents a reliable and convenient reference on molecular detection and identification of major human bacterial pathogens; an indispensable tool for upcoming and experienced medical, veterinary, and industrial laboratory scientists engaged in bacterial characterization; and an essential textbook for undergraduate and graduate students majoring in bacteriology. A comprehensive and inclusive book such as this is undoubtedly beyond an individual’s capacity. I am fortunate and honored to have a large panel of bacteriologists as chapter contributors, whose detailed knowledge and technical insights on human bacterial pathogen detection have greatly enriched this book. In addition, the professionalism and dedication of executive editor Barbara Norwitz and senior project coordinator Jill Jurgensen at CRC Press have enhanced its presentation. Finally, without the understanding and support of my family, Liling Ma, Brenda, and Cathy, the compilation of this all-encompassing volume would have not been possible.

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Editor Dongyou Liu, PhD, undertook his veterinary science education at Hunan Agricultural University, China. Upon graduation, he received an Overseas Postgraduate Scholarship from the Chinese Ministry of Education to pursue further training at the University of Melbourne, Australia, where he worked toward improved immunological diagnosis of human hydatid disease. During the past two decades, he has crisscrossed between research and clinical laboratories in Australia and the United States of America, with focuses on molecular characterization and virulence determination of microbial pathogens such as ovine footrot bacterium (Dichelobacter

nodosus), dermatophyte fungi (Trichophyton, Microsporum, and Epidermophyton), and listeriae (Listeria species). He is the senior author of more than 50 original research and review articles in various international journals and the editor of the recently released Handbook of Listeria monocytogenes, Handbook of Nucleic Acid Purification, Molecular Detection of Foodborne Pathogens, and Molecular Detection of Human Viral Pathogens, as well as the forthcoming Molecular Detection of Human Fungal Pathogens and Molecular Detection of Human Parasitic Pathogens, all of which are published by CRC Press.

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Contributors Martina Adamek Karlsruhe Institute of Technology Institute of Functional Interfaces Karlsruhe, Germany

Frank W. Austin College of Veterinary Medicine Mississippi State University Starkville, Mississippi

Dianna J. Bopp Wadsworth Center New York State Department of Health Albany, New York

Catherine C. Adley Microbiology Laboratory Department of Chemical and Environmental Sciences University of Limerick Limerick, Ireland

Mario J. Avila-Campos Department of Microbiology Institute of Biomedical Science University of São Paulo São Paulo, Brazil

Alejandra Bosch Facultad de Ciencias Exactas Centro de Investigación y Desarrollo en Fermentaciones Industriales Universidad Nacional de La Plata La Plata, Argentina

Sascha Al Dahouk Department of Internal Medicine III RWTH Aachen University Aachen, Germany Rodrigo Alonso Faculty of Pharmacy Department of Immunology, Microbiology and Parasitology University of the Basque Country Vitoria-Gasteiz, Spain Maria Silvana Alves Faculdade de Farmácia e Bioquímica Universidade Federal de Juiz de Fora Minas Gerais, Brazil

R. Aznar Department of Microbiology and Ecology University of Valencia Valencia, Spain Diprabhanu Bakshi Syngene International Bangalore, India Stephan Bathe German National Academic Foundation Bonn, Germany Harsh Vardhan Batra Defence R & D Establishment Gwalior, India

Paula Lopes Alves Instituto de Biologia Experimental e Tecnológica Oeiras, Portugal

Asim Bej Department of Biology University of Alabama at Birmingham Birmingham, Alabama

Emilio Aranda Ciencia y Tecnología de los Alimentos Escuela de Ingenierías Agrarias Badajoz, Spain

Scott C. Bell Department of Thoracic Medicine The Prince Charles Hospital Brisbane, Australia

Marco Arosio USC Microbiology and Virology A.O. Ospedali Riuniti Bergamo, Italy

María J. Benito Ciencia y Tecnología de los Alimentos Escuela de Ingenierías Agrarias Badajoz, Spain

Aurora Fernández Astorga Faculty of Pharmacy Department of Immunology, Microbiology and Parasitology University of the Basque Country Vitoria-Gasteiz, Spain

Patrick Boiron Research group on Bacterial Opportunistic Pathogens and Environment Université de Lyon Lyon, France

Javier Enrique Botero School of Dentistry Grupo de Investigación Básica y Clínica en Periodoncia y Oseointegración Universidad de Antioquia Medellín, Colombia Germán Bou Servicio Microbiología Complejo Hospitalario Universitario La Coruña La Coruña, Spain June Brown Actinomycete Reference Laboratory Bacterial Zoonoses Branch Division of Foodborne, Bacterial and Mycotic Diseases National Center for Zoonotic, VectorBorne, and Enteric Diseases Centers for Disease Control and Prevention Atlanta, Georgia Renato Corrêa Viana Casarin Department of Oral Diagnostics Division of Microbiology and Immunology Piracicaba Dental School University of Campinas Campinas, Brazil Nicholas Cassai Department of Pathology and Laboratory Medicine New York Harbor VA Medical Center Brooklyn, New York xix

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Marco Cassone Sbarro Health Research Organization College of Science and Technology Temple University Philadelphia, Pennsylvania Bianca Castiglioni Department of Veterinary Pathology, Hygiene and Public Health University of Milan Milan, Italy Rita Castro Instituto de Higiene e Medicina Tropical Universidade Nova de Lisboa Lisbon, Portugal Tsung Chain Chang Department of Medical Laboratory Science and Biotechnology College of Medicine National Cheng Kung University Tainan, Taiwan Rama Chaudhry Department of Microbiology All India Institute of Medical Sciences New Delhi, India Jianshun Chen Institute of Preventive Veterinary Medicine Zhejiang University Zhejiang, China Ameur Cherif Faculté des Sciences de Tunis Laboratoire Microorganismes et Biomolécules Actives Campus Universitaire Tunis, Tunisie Bishwanath Kumar Chourasia Department of Microbiology All India Institute of Medical Sciences New Delhi, India Henrik Christensen Faculty of Life Sciences Department of Veterinary Pathobiology Center for Applied Bioinformatics Copenhagen University Frederiksberg, Denmark

Contributors

Deirdre L. Church Departments of Pathology, Laboratory Medicine, and Medicine University of Calgary Alberta Health Services and Calgary Laboratory Services Alberta, Canada Lorenzo Ciceroni Center for Research and Evaluation of Immunobiologicals Istituto Superiore di Sanità Rome, Italy Alessandra Ciervo Department of Infectious, Parasitic and Immune-Mediated Diseases Istituto Superiore di Sanità Rome, Italy Lena Ciric Microbial Diseases UCL Eastman Dental Institute London, United Kingdom Ana Paula Vieira Colombo Department of Medical Microbiology Institute of Microbiology Federal University of Rio de Janeiro Rio de Janeiro, Brazil Andréa Vieira Colombo Institute of Biomedical Sciences State University of São Paulo Sao Paulo, Brazil Adolfo Contreras School of Dentistry Grupo de Medicina Periodontal Universidad del Valle Cali, Colombia Piet Cools Department of Clinical Biology, Immunology and Microbiology Ghent University Hospital Ghent, Belgium Juan J. Córdoba Facultad de Veterinaria Higiene de los Alimentos Universidad de Extremadura Cáceres, Spain María G. Córdoba Ciencia y Tecnología de los Alimentos Escuela de Ingenierías Agrarias Badajoz, Spain

Yvon Cormier Research Unit Laval Hospital Centre of Pneumology University of Laval Quebec, Canada Paola Cremonesi Department of Veterinary Pathology, Hygiene and Public Health University of Milan Milan, Italy Maria Teresa Barreto Crespo Instituto de Biologia Experimental e Tecnológica Oeiras, Portugal Daniele Daffonchio Dipartimento di Scienze e Tecnologie Alimentari e Microbiologiche Università degli Studi di Milano Milan, Italy Sharvari Dalal Department of Pathology SUNY Downstate Medical Center Brooklyn, New York Maria Dalamaga Laboratory of Clinical Biochemistry Attikon General University Hospital Medical School, National & Kapodistrian University of Athens Athens, Greece George N. Dalekos Department of Medicine, Medical School University of Thessaly Larissa, Greece Anupam Das Department of Microbiology All India Institute of Medical Sciences New Delhi, India Gregory A. Dasch Rickettsial Zoonoses Branch Division of Vector-Borne Diseases Centers for Disease Control and Prevention Atlanta, Georgia

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Contributors

Jonathan J. Dennis Department of Biological Sciences University of Alberta Alberta, Canada Rubens Clayton da Silva Dias Faculdade de Medicina Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Xiaoming Dong Beijing Wantai Biological Pharmacy Enterprise Beijing, China Caroline Duchaine Research Unit Laval Hospital Centre of Pneumology University of Laval Quebec, Canada Francis Dziva Division of Microbiology Institute for Animal Health Berkshire, United Kingdom P. Elizaquível Department of Microbiology and Ecology University of Valencia Valencia, Spain Marina E. Eremeeva Rickettsial Zoonoses Branch Division of Vector-Borne Diseases Centers for Disease Control and Prevention Atlanta, Georgia Emenike Ribs K. Eribe Norwegian Geotechnical Institute Oslo, Norway Klaus Eschrich Institute for Biochemistry School of Medicine University of Leipzig Leipzig, Germany Weihuan Fang Institute of Preventive Veterinary Medicine Zhejiang University Zhejiang, China

Seamus Fanning Centre for Food Safety School of Agriculture, Food Science and Veterinary Medicine Veterinary Sciences Centre University College Dublin Dublin, Ireland

John I. Glass J. Craig Venter Institute Rockville, Maryland

María Fiuza Faculty of Biology Section of Microbiology Department of Molecular Biology University of León León, Spain

Reginaldo Bruno Gonçalves Department of Oral Diagnostics Division of Microbiology and Immunology Piracicaba Dental School University of Campinas Campinas, Brazil

Maria Fredriksson-Ahomaa Ludwig-Maximilian University Munich, Germany Elerson Gaetti-Jardim Jr. Department of Oral Pathology São Paulo State University São Paulo, Brazil Ana Cristina Gales Departamento de Medicina Universidade Federal de São Paulo São Paulo, Brazil María-Luisa García-López Veterinary Faculty Department of Food Hygiene and Food Microbiology University of León León, Spain Maria del Mar Garcia-Suarez Area de Microbiologia Departamento de Biologia Funcional Instituto Universitario de Biotecnologia de Asturias Asturias, Spain José A Gil Faculty of Biology Section of Microbiology Department of Molecular Biology University of León Campus de Vegazana León, Spain Cecilia Girbau Faculty of Pharmacy Department of Immunology, Microbiology and Parasitology University of the Basque Country Vitoria-Gasteiz, Spain

Maryna Golovchenko Faculty of Science University of South Bohemia Ceské Budeˇjovice, Czech Republic

Keith Grimwood Queensland Paediatric Infectious Diseases Laboratory Royal Children’s Hospital Queensland Children’s Medical Research Institute Queensland, Australia Libor Grubhoffer Biology Centre Institute of Parasitology Academy of Sciences of the Czech Republic Ceské Budeˇjovice, Czech Republic Miguel Gueimonde Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain Saija Hallanvuo Finnish Food Safety Authority Evira National Public Health Institute Helsinki, Finland Rudy A. Hartskeerl WHO/FAO/OIE and National Collaborating Centre for Reference and Research on Leptospirosis Department KIT Biomedical Research Royal Tropical Institute Amsterdam, the Netherlands John P. Hays Department of Medical Microbiology and Infectious Diseases Rotterdam, the Netherlands

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Contributors

Alexander M. Hirschl Division of Clinical Microbiology Department of Laboratory Medicine Medical University Vienna Vienna, Austria

Adriana Jaramillo School of Dentistry Grupo de Medicina Periodontal Universidad del Valle Cali, Colombia

Jeffrey Hoorfar National Food Institute Technical University of Denmark Søborg, Denmark

Mathilde H. Josefsen National Food Institute Technical University of Denmark Søborg, Denmark

Kurt Houf Faculty of Veterinary Medicine Department of Veterinary Public Health and Food Safety Ghent University Ghent, Belgium

Estelle Jumas-Bilak Universitaire de Montpellier Hôpital Arnaud de Villeneuve Laboratoire de Bactériologie Montpellier, France

Satoru Inagaki Department of Microbiology Oral Health Science Center Tokyo Dental College Chiba, Japan

Bernhard Kaltenboeck Molecular Diagnostics Laboratory Department of Pathobiology College of Veterinary Medicine Auburn University Auburn, Alabama

Kazuyuki Ishihara Department of Microbiology Oral Health Science Center Tokyo Dental College Chiba, Japan Nahed Ismail Department of Pathology Meharry Medical College Nashville, Tennessee Carol Iversen Centre for Food Safety School of Agriculture Food Science and Veterinary Medicine Veterinary Sciences Centre University College Dublin Dublin, Ireland Jari Jalava Department of Infectious Disease Surveillance and Control National Institute for Health and Welfare Turku, Finland Judith Jansen Medical Faculty Institute of Immunology RWTH Aachen University Aachen, Germany

Mireille M. Kattar Department of Laboratory Medicine and Pathology University of Alberta Hospital Alberta, Canada Timothy J. Kidd Queensland Paediatric Infectious Diseases Laboratory Royal Children’s Hospital Queensland Children’s Medical Research Institute Queensland, Australia Rickard Knutsson Swedish National Veterinary Institute Uppsala, Sweden Eija Könönen University of Turku Institute for Dentistry Turku, Finland Hannu Korkeala Faculty of Veterinary Medicine Department of Food and Environmental Hygiene University of Helsinki Helsinki, Finland

Kiersten J. Kugeler Centers for Disease Control and Prevention National Center for Zoonotic VectorBorne and Enteric Diseases Division of Vector-Borne Infectious Diseases, Bacterial Diseases Branch Fort Collins, Colorado Susanne Thisted Lambertz Research and Development Department National Food Administration Uppsala, Sweden K.A. Lampel Food and Drug Administration Division of Microbiology College Park, Maryland Brent A. Lasker Actinomycete Reference Laboratory Bacterial Zoonoses Branch Division of Foodborne, Bacterial and Mycotic Diseases National Center for Zoonotic, VectorBorne, and Enteric Diseases Centers for Disease Control and Prevention Atlanta, Georgia Susanna K.P. Lau Department of Microbiology The University of Hong Kong Hong Kong Kevin B. Laupland Departments of Medicine, Critical Care Medicine, Community Health Sciences, and Pathology and Laboratory Medicine, and Centre for Antimicrobial Resistance University of Calgary, Alberta Health Services, and Calgary Laboratory Services Alberta, Canada Frédéric Laurent Faculté de Médecine Laennec Lyon, France Marc Lehmann SIRS-Lab GmbH Jena, Germany Angelika Lehner Vetsuisse Faculty Institute for Food Safety and Hygiene University of Zurich Zurich, Switzerland

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Contributors

Manuel L. Lemos Department of Microbiology and Parasitology Institute of Aquaculture University of Santiago de Compostela Santiago de Compostela, Galicia, Spain

Angel Manteca Area de Microbiologia Departamento de Biologia Funcional Instituto Universitario de Biotecnologia de Asturias Universidad de Oviedo Asturias, Spain

Michal Letek Faculty of Biology Section of Microbiology Department of Molecular Biology University of León Campus de Vegazana León, Spain

Hélène Marchandin Faculté de Pharmacie Université Montpellier 1 Laboratoire de Bactériologie-VirologieContrôle Microbiologique Montpellier, France

Dongyou Liu BioSecurity Quality Assurance Program Royal College of Pathologists of Australasia New South Wales, Australia

Abelardo Margolles Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain

Charlotta Löfström National Food Institute Technical University of Denmark Søborg, Denmark José Maria Luengo Department of Molecular Biology University of León León, Spain Karlene H. Lynch Department of Biological Sciences University of Alberta Alberta, Canada Athanasios Makristathis Division of Clinical Microbiology Department of Laboratory Medicine Medical University Vienna Vienna, Austria Fabiola Mancini Department of Infectious, Parasitic and Immune-Mediated Diseases Istituto Superiore di Sanità Rome, Italy Chaysavanh Manichanh Digestive System Research Unit University Hospital Vall d’Hebron Bioinformatics and Genomics Program Center for Genomic Regulation Barcelona, Spain

Luis M. Mateos Faculty of Biology Section of Microbiology Department of Molecular Biology University of León Campus de Vegazana León, Spain Johanna Maukonen VTT Technical Research Centre of Finland Espoo, Finland Jere W. McBride Department of Pathology University of Texas Medical Branch at Galveston Galveston, Texas Andrew McDowell Centre of Infection & Immunity School of Medicine, Dentistry and Biomedical Sciences Queen’s University Belfast, Northern Ireland Francisco J. Mendez Area de Microbiologia Departamento de Biologia Funcional Instituto Universitario de Biotecnologia de Asturias Universidad de Oviedo Asturias, Spain

Kåre Mølbak Statens Serum Institut Copenhagen, Denmark Jens Kjølseth Møller Department of Clinical Microbiology Institute of Regional Health Services Research University of Southern Denmark Odense, Denmark Beatriz Meurer Moreira Instituto de Microbiologia Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Paolo Moroni Quality Milk Production Services Cornell University Ithaca, New York Benjamin D. Moser Actinomycete Reference Laboratory Bacterial Zoonoses Branch Division of Foodborne, Bacterial and Mycotic Diseases National Center for Zoonotic, VectorBorne, and Enteric Diseases Centers for Disease Control and Prevention Atlanta, Georgia Dipalok Mukherjee Biological Sciences Mississippi State University Starkville, Mississippi Soma Mukherjee Palli Shikha Bhavan Sriniketan, Birbhum West Bengal, India Kimberlee A. Musser Wadsworth Center New York State Department of Health Albany, New York Beth Mutai Global Emerging Infections Surveillance System U.S. Army Medical Research Unit Kenya Medical Research Institute Narombi, Kenya

xxiv

Germán Naharro Department of Animal Health University of León León, Spain Futoshi Nakazawa Department of Oral Microbiology School of Dentistry, Health Sciences University of Hokkaido Hokkaido, Japan

Contributors

Carlos R. Osorio Department of Microbiology and Parasitology Institute of Aquaculture University of Santiago de Compostela Compostela, Spain Jie Ouyang Department of Pathology SUNY Downstate Medical Center Brooklyn, New York

Elizabeth J. Nazarian Wadsworth Center New York State Department of Health Albany, New York

Vanya Paralanov J. Craig Venter Institute Rockville, Maryland

Heinrich Neubauer Friedrich-Loeffler-Institut Institute of Bacterial Infections and Zoonoses Jena, Germany

Sheila Patrick Centre of Infection & Immunity School of Medicine, Dentistry and Biomedical Sciences Queen’s University Belfast, Northern Ireland

Juan P. Olano Department of Pathology University of Texas Medical Branch at Galveston Galveston, Texas

Biswajit Paul Escorts Heart Institute and Research Centre New Delhi, India

James H. Oliver, Jr. Georgia Southern University The James H. Oliver, Jr. Institute of Arthropodology and Parasitology Statesboro, Georgia Ingar Olsen Faculty of Dentistry Institute of Oral Biology University of Oslo Oslo, Norway Katharina E.P. Olsen Statens Serum Institut Copenhagen, Denmark

Debarati Paul College of Veterinary Medicine Mississippi State University Starkville, Mississippi Debosmita Paul Jamia Millia Islamia New Delhi, India Ana Isabel Pelaez Area de Microbiologia Departamento de Biologia Funcional Instituto Universitario de Biotecnologia de Asturias Universidad de Oviedo Asturias, Spain

Efrén Ordoñez Faculty of Biology Section of Microbiology Department of Molecular Biology University of León León, Spain

Claire Pelletier Laboratoire Départemental d’Analyses de Saône-et-Loire Mâcon, France

Andrés Otero Veterinary Faculty Department of Food Hygiene and Food Microbiology University of León León, Spain

J. Tony Pembroke Molecular and Structural Biochemistry Laboratory Department of Chemical and Environmental Sciences University of Limerick Limerick, Ireland

Filomena Martins Pereira Instituto de Higiene e Medicina Tropical Universidade Nova de Lisboa Lisbon, Portugal Jeannine M. Petersen Centers for Disease Control and Prevention National Center for Zoonotic, VectorBorne and Enteric Diseases Division of Vector-Borne Infectious Diseases, Bacterial Diseases Branch Fort Collins, Colorado Efthimia Petinaki Department of Microbiology University of Thessaly Medical School Larissa, Greece Wolfgang Pfister Institute for Medical Microbiology University Hospital of Jena Jena, Germany Matthew R. Pincus Department of Pathology SUNY Downstate Medical Center Brooklyn, New York Giuliano Pisoni Department of Veterinary Pathology, Hygiene and Public Health University of Milan Milan, Italy Brunella Posteraro Institute of Microbiology Università Cattolica del Sacro Cuore Rome, Italy Patrizia Posteraro Laboratory of Clinical Pathology and Microbiology Ospedale San Carlo Rome, Italy Noura Raddadi Dipartimento di Scienze e Tecnologie Alimentari e Microbiologiche Università degli Studi di Milano Milan, Italy Peter Rådström Department of Applied Microbiology Lund University Lund, Sweden

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Contributors

Annibale Raglio USC Microbiology and Virology A.O. Ospedali Riuniti Bergamo, Italy

Patricia Ruas-Madiedo Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain

Borja Sánchez Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain

D. Madhusudan Reddy Palamuru University Andhra Pradesh, India

Pedro Rubio Department of Animal Health University of León León, Spain

Clara G. de los Reyes-Gavilán Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain

Nataliia Rudenko Biology Centre Institute of Parasitology Academy of Sciences of the Czech Republic Cˇ eské Budeˇjovice, Czech Republic

Ester Sánchez Microbial Ecophysiology and Nutrition Research Group Institute of Agrochemistry and Food Technology Spanish Council for Scientific Research Valencia, Spain

Lothar Rink Institute of Immunology Medical Faculty RWTH Aachen University Aachen, Germany

Stefan Rupf Clinic of Operative Dentistry Periodontology and Preventive Dentistry Saarland University Hospital Homburg/Saar, Germany

Ignasi Roca Department of Clinical Microbiology School of Medicine University of Barcelona Barcelona, Spain

Michael P. Ryan Microbiology Laboratory Department of Chemical and Environmental Sciences University of Limerick Limerick, Ireland

Derren Ready Eastman Dental Hospital UCLH NHS Foundation Trust London, United Kingdom

Camilla Rodrigues P D Hinduja National Hospital & Medical Research Centre Mumbai, India Mar Rodríguez Facultad de Veterinaria Higiene de los Alimentos Universidad de Extremadura Cáceres, Spain Veronica Rodriguez-Nava Research group on “Bacterial Opportunistic Pathogens and Environment” Université de Lyon Lyon, France Antoni Róz·alski Department of Immunobiology of Bacteria Institute of Microbiology and Immunology University of Łódz´ Łódz´, Poland

Maria Saarela VTT Technical Research Centre of Finland Espoo, Finland Konrad Sachse Friedrich-Loeffler-Institut (Federal Research Institute for Animal Health) Institute of Molecular Pathogenesis Jena, Germany Atsushi Saito Oral Health Science Center Tokyo Dental College Chiba, Japan Daniel Saito Department of Oral Diagnostics Division of Microbiology and Immunology Piracicaba Dental School University of Campinas Campinas, Brazil

G. Sánchez Department of Biotecnology Institute of Agrochemistry and Food Technology Valencia, Spain Maurizio Sanguinetti Institute of Microbiology Università Cattolica del Sacro Cuore Rome, Italy Guido Lopes dos Santos Santiago Department of Clinical Biology, Immunology and Microbiology Ghent University Hospital Ghent, Belgium Jesús A. Santos Veterinary Faculty Department of Food Hygiene and Food Microbiology University of León León, Spain Yolanda Sanz Microbial Ecophysiology and Nutrition Research Group Institute of Agrochemistry and Food Technology Spanish Council for Scientific Research Valencia, Spain Roland P.H. Schmitz SIRS-Lab GmbH Jena, Germany

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Contributors

K.R. Schneider Food Science and Human Nutrition Department University of Florida Gainesville, Florida

Renata Martins do Souto Department of Medical Microbiology Institute of Microbiology Federal University of Rio de Janeiro Rio de Janeiro, Brazil

María Mar Tomás Servicio Microbiología Complejo Hospitalario Universitario La Coruña La Coruña, Spain

Bo Segerman Swedish National Veterinary Institute Uppsala, Sweden

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

Herbert Tomaso Friedrich-Loeffler-Institut Institute of Bacterial Infections and Zoonoses Jena, Germany

Teresa Semedo-Lemsaddek Faculdade de Ciências Universidade de Lisboa Center for Biodiversity Functional and Integrative Genomics Edifício ICAT, Campus da FCUL Lisbon, Portugal Diego Omar Serra Facultad de Ciencias Exactas Centro de Investigación y Desarrollo en Fermentaciones Industriales Universidad Nacional de La Plata La Plata, Argentina Nidhi Sharma Department of Microbiology All India Institute of Medical Sciences New Delhi, India Shubhada Shenai P D Hinduja National Hospital & Medical Research Centre Mumbai, India Han Shiong Siah TPP, Lambells Lagoon Northern Territory, Australia Shoo Peng Siah Human Genetic Signatures New South Wales, Australia Maria R.L. Simionato Department of Microbiology Institute of Biomedical Science University of São Paulo São Paulo, Brazil Mikael Skurnik University of Helsinki and Helsinki University Central Hospital Laboratory Diagnostics Helsinki, Finland

Paola Stefanelli Department of Infectious, Parasitic and Immune-Mediated Diseases Istituto Superiore di Sanità Rome, Italy Roger Stephan Vetsuisse Faculty Institute for Food Safety and Hygiene University of Zurich Zurich, Switzerland L.K. Strawn Department of Food Science Cornell University Ithaca, New York Yi-Wei Tang Departments of Pathology and Medicine Vanderbilt University School of Medicine Nashville, Tennessee Marleen Temmerman Department of Obstetrics and Gynaecology Ghent University Ghent, Belgium Rogério Tenreiro Faculdade de Ciências Universidade de Lisboa Center for Biodiversity Functional and Integrative Genomics Edifício ICAT, Campus da FCUL Lisbon, Portugal Corinne Teyssier Faculté de Pharmacie, BP 14491 Université Montpellier 1 Montpellier, France

Martha E. Trujillo Departmento de Microbiología y Genética Edificio Departamental Universidad de Salamanca Salamanca, Spain Sheng Kai Tung Department of Medical Laboratory Science and Biotechnology College of Medicine National Cheng Kung University Taiwan Mario Vaneechoutte Department of Clinical Biology, Immunology and Microbiology Ghent University Hospital Ghent, Belgium A.C.M. Veloo Department of Medical Microbiology University Medical Center Groningen University of Groningen Groningen, the Netherlands Suzanne J.C. Verhaegh Department of Medical Microbiology and Infectious Diseases, Erasmus MC Rotterdam, the Netherlands Rita Verhelst Department of Clinical Biology, Immunology and Microbiology Ghent University Hospital Ghent, Belgium Hans Verstraelen Department of Obstetrics and Gynaecology Ghent University Hospital Ghent, Belgium

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Contributors

Jordi Vila Department of Clinical Microbiology Hospital Clinic University of Barcelona School of Medicine Barcelona, Spain

Chengming Wang Molecular Diagnostics Laboratory Department of Pathobiology College of Veterinary Medicine Auburn University Auburn, Alabama

Almudena F. Villadangos Section of Microbiology Department of Molecular Biology Faculty of Biology University of León León, Spain

B.R. Warren Research, Quality and   Innovation ConAgra Foods, Inc. Omaha, Nebraska

Georgia Vrioni Department of Microbiology National & Kapodistrian University of Athens Medical School Athens, Greece Ken B. Waites Department of Pathology University of Alabama at Birmingham Birmingham, Alabama

David M. Whiley Queensland Paediatric Infectious Diseases Laboratory Royal Children’s Hospital Queensland Children’s Medical Research Institute Queensland, Australia Patrick C.Y. Woo Department of Microbiology The University of Hong Kong Hong Kong

Li Xiao Department of Pathology University of Alabama at Birmingham Birmingham, Alabama Osvaldo Miguel Yantorno Facultad de Ciencias Exactas Centro de Investigación y Desarrollo en Fermentaciones Industriales Universidad Nacional de La Plata La Plata, Argentina Ting Zhang College of Veterinary Medicine Mississippi State University Starkville, Mississippi

1 Introductory Remarks Dongyou Liu CONTENTS 1.1 Preamble............................................................................................................................................................................... 1 1.2 Bacterial Characteristics....................................................................................................................................................... 2 1.2.1 Classification............................................................................................................................................................. 2 1.2.2 Morphology.............................................................................................................................................................. 2 1.2.3 Biology...................................................................................................................................................................... 4 1.2.4 Genetics.................................................................................................................................................................... 5 1.2.5 Ecological and Medical Importance......................................................................................................................... 5 1.3 Bacterial Identification......................................................................................................................................................... 6 1.3.1 Current Diagnostic Approaches............................................................................................................................... 6 1.3.2 Performance Parameters........................................................................................................................................... 7 1.3.3 Result Interpretation................................................................................................................................................. 7 1.3.4 Standardization, Quality Control, and Assurance.................................................................................................... 8 1.4 Conclusion............................................................................................................................................................................ 9 References...................................................................................................................................................................................... 9

1.1  PREAMBLE Bacteria (singular, bacterium) are small unicellular organisms that are classified taxonomically in the domain Bacteria (or Eubacteria), the kingdom Prokaryotae (or Prokaryota or Monera). The only other domain in the kingdom Prokaryotae covers Archaea (or Archaeobacteria for “ancient bacteria”). With sizes ranging from 10 −7 to 10 −4 mm, prokaryotes are bigger than viruses (10 −8–10 −6 mm), but smaller than eukaryotes (10 −5–103 mm). While both bacteria and archaea may have evolved independently from an ancient common ancestor, eukaryotes may have arisen from ancient bacteria entering into endosymbiotic associations with the ancestors of eukaryotic cells (possibly related to the archaea) to form either mitochondria or hydrogenosomes. A subsequent independent engulfment by some mitochondria-containing eukaryotes of cyanobacterial-like organisms may have led to the formation of chloroplasts in algae and plants. In contrast to the organisms in the eukaryotic kingdoms Protista, Fungi, Plantae, and Animalia, those in the kingdom Prokaryotae lack nuclear membrane (with their DNA usually in a loop or coil), contain few independent membrane-bounded cytoplasmic organelles (e.g., vacuole, endoplasmic reticulum, Golgi apparatus, and mitochondria) apart from chromosome and ribosome, have no unique structures in their plasma membrane and cell wall, and do not undergo endocystosis and exocytosis. In other words, whereas eukaryotic chromosome resides within a membrane-delineated nucleus, bacterial chromosome is located inside the

bacterial cytoplasm. This entails that all cellular events (e.g., translational and transcriptional processes, and interaction of chromosome with other cytoplasmic structures) in prokaryotes occur in the same compartment. Furthermore, while eukaryotic chromosome is packed with histones to form linear chromatin, bacterial chromosome assumes a highly compact supercoiled structure in circular form (and rarely in linear form). Although archaea are similar to bacteria in most aspects of cell structure and metabolism, they differ from bacteria in that being extremophiles, they can live in extreme environments where no other life forms exist. This may be due to the unique structure in archaeal lipids in which the stereochemistry of the glycerol is the reverse of that found in bacteria and eukaryotes, possibly the result of an adaptation on the part of archaea to hyperthermophily. In addition, the archaeal cell wall does not contain muramic acid, which is commonly present in bacteria. The archaeal RNA polymerase core is composed of ten subunits in comparison with four subunits in bacteria. Besides possessing distinct tRNA and rRNA genes, archaea uses eukaryotic-like initiation and elongation factors in protein translation, and their transcription involves TATA-binding proteins and TFIIB as in eukaryotes. Bacteria are ubiquitously distributed in virtually every habitat on earth, and are abundantly present in soil, fresh water, plants, and animals. With an estimated number of 5  nonillion (5 × 1030), bacteria form much of the world’s biomass. Bacteria play an essential role in chemical cycles, 1

2

Molecular Detection of Human Bacterial Pathogens

environmental maintenance, food production, and human wellbeing. However, some bacteria are pathogenic and capable of causing infectious diseases in humans, animals, and plants. Cholera, syphilis, anthrax, leprosy, bubonic plague, and tuberculosis are some of the examples of the deadly human diseases that are attributable to bacteria. Correct identification and detection of bacterial pathogens is not only fundamental to the study of these microorganisms but also critical to the control and prevention of the diseases they cause.

1.2  BACTERIAL CHARACTERISTICS

microbia, Thermotogae, and Verrcomicrobia), whereas  the domain Archaea (Archaeobacteria) is separated into two phyla (Crenarchaeota and Euryarchaeota). Among the 26 phyla in the domain Bacteria, Proteobacteria, and Firmicutes contain the largest numbers of genera and species followed by Cyanobacteria, Bacteroidetes, Spirochaetes, and Flavobacteria. Bacteria from other phyla are comparatively rare, from which fewer genera and species have been described. Most of human pathogenic bacteria are found in the phyla Actinobacteria, Bacteroidetes, Chlamydiae, Firmicutes, Fusobacteria, Proteobacteria, Spirochaetes, and Tenericutes (Table 1.1).

1.2.1  Classification

1.2.2  Morphology

Bacteria are classified on the basis of their differences in morphology (e.g., rod, cocci, spirilla, and filament), cell  wall structure (e.g., gram-negative and gram-positive), growth characteristics (e.g., aerobic and anaerobic), biochemical properties (e.g., fatty acids), and genetic features (e.g., 16S and 23S rRNA). Currently, the domain Bacteria (Eubacteria) is divided into 26 phyla (Acidobacteria, Actinobacteria, Aquificae, Bacteroidetes, Chlamydiae, Chlorobi, Chloroflexi, Chrysiogenetes, Cyanobacteria, Deferribacteres, Deinococcus-Thermus, Dictyoglomi, Fibro­ bacteres, Firmicutes, Fusobacteria, Gemmatimonadetes, Lentisphaerae, Nitrospira, Planctomycetes, Proteobacteria, Spirochaetes, Tenericutes, Thermodesulfobacteria, Thermo­

Bacteria usually measure from 0.2 to 2.0 μm in width and 2–8  μm in length, and are 10 times smaller than eukaryotic cells. On one extreme, a few bacterial species (e.g., Thiomargarita namibiensis, and Epulopiscium fishelsoni) measure up to half a mm long and are visible to the naked eye. On the other extreme, the smallest bacteria in the genus Mycoplasma are only 0.3 μm in size, which are as small as the largest viruses. Bacteria typically assume four distinctive forms: rodlike bacilli, spherical cocci, spiral bacteria (also called spirilla), and filamentous bacteria. Occasionally, a small number of bacterial species may appear tetrahedral or cuboidal in shape. While many bacterial species exist as

TABLE 1.1 Classification and Characteristics of Major Human Bacterial Pathogens Phylum

Class

Actinobacteria

Actinobacteria

Firmicutes

Bacilli Clostridia

Tenericutes

Mollicutes

Bacteroidetes

Bacteroidia

Chlamydiae Fusobacteria Proteobacteria

Flavobacteria Chlamydiae Fusobacteria Alphaproteobacteria Betaproteobacteria Gammaproteobacteria

Epsilonproteobacteria Spirochaetes

Spirochaetes

Brief Description Gram-positive bacteria with high G + C ratio; classification may be assisted through analysis of ferric uptake regulator (fur) and glutamine synthetase Gram-positive cocci or rods with low G + C ratio; presence of cell wall Gram-positive cocci or rods with low G + C ratio; presence of cell wall; some species (e.g., Veillonella) are gram-negative Small bacteria (0.2–0.3 μm in size) with low G + C ratio; absence of cell wall (outer membrane) Gram-negative, anaerobic bacteria; opportunistic pathogens Gram-negative, anaerobic bacteria; opportunistic pathogens Gram-negative bacteria; obligate intracellular pathogens Gram-negative, filamentous, anaerobic bacteria Gram-negative, phototrophic bacteria, with symbiotic properties Gram-negative, aerobic or facultative bacteria some of which are chemolithotrophs, or phototrophs Gram-negative, facultatively or obligately anaerobic bacteria, some of which are highly pathogenic Gram-negative, curved to spirilloid bacteria, inhabiting digestive tract Gram-negative, long, helically coiled (spiral-shaped), chemoheterotrophic, anaerobic bacteria, with length-wise flagella

Notable Human Pathogens Actinomyces, Corynebacterium, Mycobacterium, Nocardia Bacillus, Enterococcus, Listeria, Staphylococcus, Streptococcus Clostridium, Eubacterium, Peptostreptococcus Mycoplasma, Ureaplasma Bacteroides, Porphyromonas, Prevotella Elizabethkingia, Flavobacterium Chlamydia, Chlamydophila Fusobacterium, Leptotrichia Bartonella, Brucella Bordetella, Burkholderia, Neisseria Aeromonas, Klebsiella, Pseudomonas, Salmonella, Vibrio, Yersinia Arcobacter, Campylobacter, Helicobacter Borrelia, Leptospira, Treponema

Introductory Remarks

single cells, others present characteristic patterns such as diploids (pairs), chains, and clusters (“bunch of grapes”). In addition, some bacteria may be elongated to form filaments, which are often surrounded by a sheath containing many individual cells. The elaborated, branched filaments formed by Nocardia may even resemble fungal mycelia in appearance. Frequently, bacteria use quorum sensing to detect surrounding cells, migrate toward each other, and attach to solid surfaces to form dense aggregations called biofilms (bacterial mats, or fruiting bodies), which may measure a few micrometers in thickness to up to half a meter in depth, and which comprise multiple species of bacteria, archaea, and protists (numbering approximately 100,000 cells). The formation of biofilms protects bacteria from host defense mechanisms and antibiotic therapy, contributing to chronic bacterial infections and infections relating to implanted medical devices. Structurally, a bacterial cell is surrounded by a rigid layer (cell wall) that is located externally to the lipid membrane. The cell wall provides structural support and protection, and acts as a filtering mechanism. In addition to prokaryotae, fungi and plantae also possess a cell wall, but animalia and most protista do not. While the bacterial cell wall is made up of peptidoglycan (also called murein, which in turn is composed of polysaccharide chain cross-linked by peptides containing d-amino acids), the archaeal cell wall consists of surface layer proteins (also known as S-layer), pseudopeptidoglycan (pseudomurein), and polysaccharides. By contrast, the fungal cell wall includes chitin, the algal cell wall has glycoprotein and polysaccharides, and the plant cell wall often incorporates cellulose and proteins such as extensins. Based on the ability of bacterial cell wall to retain Gram stain (consisting of crystal violet as primary stain and Gram’s iodine and basic fuchsin as subsequent stain), bacteria are divided into gram-positive and gram-negative categories. The gram-positive bacterial cell wall is composed of several layers of peptidoglycan (which is responsible for retaining the crystal violet dyes during the Gram staining procedure, leading to its purple color) surrounded by a second lipid membrane containing lipopolysaccharides and lipoproteins. Located outside of cytoplasmic membrane, peptidoglycan is a large polymer (formed by poly-N-acetylglucosamine and N-acetylemuramic acid) that contributes to the structural integrity of the bacterial cell wall in addition to countering the osmotic pressure of the cytoplasm. Peptidoglycan is predominant in the cell walls of high and low percentage G + C gram-positive organisms (e.g., actinobacteria and firmicutes). Also imbedded in the gram-positive cell wall are teichoic acids, some of which are lipid linked to form lipoteichoic acids. On the other hand, the gram-negative cell wall has a thin peptidoglycan layer adjacent to the cytoplasmic membrane that contributes to its inability to retain the crystal violet stain upon decolonization with ethanol during the Gram staining procedure (leading to its red or pink color after restaining with basic fuchsin). Apart from the thin peptidoglycan layer, the gram-negative cell wall also

3

has an outer membrane that is formed by phospholipids and lipopolysaccharides. Within the gram-positive bacterial category, there is another distinct group of bacteria (i.e., acid-fast bacteria such as Mycobacterium and Nocardia) that can resist decolorization with an acid-alcohol mixture during the acid-fast (or Ziehl–Neelsen) staining procedure and retain the initial dye carbol fuchsin and appear red. The acid-fast cell wall of Mycobacterium includes a large amount of glycolipids, especially mycolic acids that make up approximately 60% of the acid-fast cell wall in addition to a thin, inner-layer peptidoglycan. The presence of the mycolic acids and other glycolipids impede the entry of chemicals, causing the organisms to grow slowly and be more resistant to chemical agents and lysosomal components of phagocytes than most bacteria. Whereas a vast majority of bacteria possess the gramnegative cell wall, the firmicutes and actinobacteria (previously known as the low percentage G + C and high percentage G + C gram-positive bacteria, respectively) have the gram-positive structure, and the tenericutes (e.g., the genus Mycoplasma) are devoid of a cell wall in spite of their similarity in G + C ratio to the firmicutes. The differences in the cell wall often determine the susceptibility and resistance of bacteria to antibiotics and other therapeutic reagents. Given that Mycoplasma species lack a cell wall, they are unaffected by such commonly used antibiotics such as penicillin and streptomycin that target cell wall synthesis. With their small size (0.3 μm), Mycoplasma species are often identified as a source of contaminating infection in the cell culture (where penicillin and streptomycin are incorporated in the culture media), causing retarded growth of cultured cell lines. The cell wall of bacteria forms part of pathogen-associated molecular patterns (or PAMPs), which are recognized by pattern-recognition receptors (or PRRs) in mammalian hosts to initiate and promote innate and adaptive immune defenses against invading bacteria. Several recognizable extracellular structures are present in bacteria. These include flagella, pili, and fimbriae, which protrude from bacterial cell wall and are involved in bacterial twitching movement as well as interaction with one another and other organisms. Bacterial flagellum (measuring 20 nm in diameter and up to 20 μm in length) is a long, whip-like, and helical projection made up of repeating flagellin protein. The numbers and arrangements of flagella vary among bacterial genera and species. Monotrichous bacteria have a single flagellum, amphitrichous bacteria contain a single flagellum on each of cell poles, lophotrichous bacteria include multiple flagella that are located at one cell pole, and peritrichous bacteria have multiple flagella that are situated at several locations. Flagella in bacteria are powered by a flow of H+ ions (sometimes Na+ ions), and those in archaea are powered by adenosine 5'-triphoshate (ATP). Despite having a similar appearance, eukaryotic flagella (called cilia or undulipodia) differ from prokaryotic flagella in both structure and evolutionary origin. A eukaryotic flagellum is a bundle of nine fused pairs of microtubule doublets surrounding two single microtubules. Eukaryotic flagella are often arranged

4

en masse at the surface of a stationary cell anchored within an organ, lashing back and forth and serving to move fluids along mucous membranes such as trachea. In addition, some eukaryotic cells (e.g., rod photoreceptor cells of eye, olfactory receptor cells of nose, and kinocilium in cochlea of ear) have immotile flagella that function as sensation and signal transduction devises. Pilus and fimbria are proteinaceous, hair- or thread-like appendages in bacteria (particularly of gram-negative category) that are much shorter and thinner than flagellum. Bacteria have up to ten pili (typically 6–7 nm in diameter) whose main function is to connect the bacterium to another of the same or a different species to enable transfer of plasmids between the bacteria (i.e., conjugation). A fimbria (measuring 2–10 nm in diameter and up to several μm in length) is shorter than pilus. A bacterium possesses as many as 1000 fimbriae, which are deployed to attach to surface of another bacterium (to form a biofilm) or host cell (to facilitate invasion). Many pilin proteins are characteristic among bacterial species and subgroups, which have been exploited as targets for serological typing of bacteria (serotypes or serovars). Many bacteria produce capsules or slime layers around their cells, which can protect cells from engulfment by eukaryotic cells (e.g., macrophages), act as antigens for cell recognition, and aid attachment to surfaces and the formation of biofilms. In addition, some gram-positive bacteria (e.g., Bacillus, Clostridium, and Anaerobacter) can form highly resistant, dormant structures called endospores, which contain a central core of cytoplasm with DNA and ribosomes surrounded by a cortex layer and protected by an impermeable and rigid coat. Endospores can survive extreme physical and chemical stresses (e.g., UV lights, γ-radiation, detergents and disinfectants, heat, pressure, and desiccation), and may remain viable for millions of years. Endospore-forming bacteria (e.g., Bacillus anthrax and Clostridium tetanus) are also capable of causing disease. Underneath the lipid membrane is the cytoplasm, which is composed of nutrients (or nutrient storage granules such as glycogen, polyphosphate, sulfur, or polyhydroxyalkanoates), proteins, and other essential components. There is a notable absence of membrane-bound organelles (with the exception of chromosome and ribosome) in the bacterial cytoplasm, although certain subcellular compartments (prokaryotic cytoskeleton), such as carboxysome-containing polyhedral protein shells, have been detected. These polyhedral organelles compartmentalize bacterial metabolism, similar to the function performed by the membrane-bound organelles in eukaryotes. The bacterial chromosome consists of a single circular DNA molecule that is situated together with associated proteins and RNA in an irregularly shaped body called the nucleoid. The bacterial ribosomes are responsible for production of proteins.

1.2.3  Biology Bacteria utilize many metabolic pathways (e.g., glycolysis, electron transport chains, chemiosmosis, cellular respiration,

Molecular Detection of Human Bacterial Pathogens

and photosynthesis) and thus virtually all carbon or energy supplies for their maintenance and growth. They are easily grown using either solid or liquid media (e.g., Luria Bertani broth). Solid growth media (e.g., agar plates) are useful for isolation of pure cultures of a bacterial strain, and liquid growth media are employed to generate bulk quantities of bacterial cells. In addition, selective media (containing specific nutrients and antibiotics) assist the isolation and identification of specific bacterial organisms. As single-celled organisms, prokaryotes reproduce by asexual binary fission, which begins with DNA replication within the cell until the entire prokaryotic DNA is duplicated. The two chromosomes then separate as the cell grows and the cell membrane invaginates, splitting the cell into two daughter cells. This reproductive process is highly efficient and leads to exponential growth of bacteria. In fact, under optimal growth conditions, Escherichia coli cells can double every 20 min. Because bacteria are able to multiply rapidly with minimal nutritional requirements, they are abundant in virtually every habitat on earth. In soil, bacteria live by degrading organic compounds and assist in soil formation. In aquatic environments such as ponds, streams, lakes, rivers, seas, and oceans, bacteria such as cyanobacteria (sometimes called blue-green algae because of their color) utilize their chlorophylls to capture energy from the sunlight. In the depths of the sea, bacteria obtain energy from oxidizing or reducing naturally occurring sulfur compounds. In humans and animals, bacteria are found in large numbers on the skin, the respiratory and digestive tracts, and other parts of the body, constituting a normal microbiota in an essentially symbiotic relationship with mutual benefits. Although the vast majority of bacteria are harmless and sometimes even beneficial to their hosts, a few have the capacity to take advantage of temporary weakness in the host (e.g., injury and/or impaired immune function) to cause diseases of varying severity. In a high-nutrient environment, the growth cycle of bacteria usually undergoes three phases. The first phase (the lag phase) is a period of slow growth with the bacterial cells adapting to the high-nutrient environment and preparing for fast growth. In the lag phase, the cell replicates its DNA and makes all the other molecules (e.g., ribosomes, membrane transport proteins) needed for the new cell. The second phase (the logarithmic phase or “log” phase, also known as the exponential phase) occurs when DNA replication stops, and is characterized by rapid cell division and exponential growth. The rate at which cells grow during this phase is known as the growth rate, and the time it takes the cells to double is known as the generation time. During the log phase, nutrients are metabolized at maximum speed until one of the nutrients is depleted, which poses a negative impact on growth. The final phase (the stationary phase) results from the depletion of nutrients. During the stationary phase, the cells decrease their metabolic activity and consume nonessential cellular proteins. As a transition from rapid growth to a stress response state, there is heightened expression of genes involved in DNA repair, antioxidant metabolism,

5

Introductory Remarks

and nutrient transport. Although the entire cycle of bacterial growth takes about an hour, a rapidly growing bacterial cell carries out multiple rounds of replication simultaneously, which helps to shorten the doubling time for most bacteria to about 20 min.

1.2.4  Genetics Bacteria have a single circular chromosome that ranges in size from only 160,000 bp (base pairs) (e.g., Candidatus Carsonella ruddii) to 12,200,000 bp (e.g., Sorangium cellulosum). However, Borrelia burgdorferi, the causal agent for Lyme disease, contains a single linear chromosome. In addition, bacteria may possess small extrachromosomal DNA called plasmid, which ranges from 1 to 400 kb in size and comprises genes or gene cassettes for antibiotic resistance or virulence factors. As plasmids have at least an origin of replication (or ori)—a starting point for DNA replication— they are capable of autonomous replication independent of the chromosomal DNA. A plasmid that integrates into the chromosomal DNA is called episome, which permits its duplication with every cell division of the host. Some viruses (bacteriophages or phages) may also exist in bacteria, with some merely infecting and lysing their host bacteria, while others inserting into the bacterial chromosome. Phages are usually made up of a nucleic acid core (e.g., ssRNA, dsRNA, ssDNA, or dsDNA measuring 5–500 bp in length) with an outer protein hull. A phage containing particular genes may contribute to its host’s phenotype, as illustrated by the evolution of Escherichia coli O157:H7 and Clostridium botulinum, which are converted from harmless ancestral bacteria into lethal pathogens through the integration of phages harboring toxin genes. Being the key component of the ribosome, ribosomal RNA molecules (rRNA) consists of two complex folded subunits of differing sizes (small and large), whose main functions are to provide a mechanism for decoding messenger RNA (mRNA) into amino acids (at center of small ribosomal subunit) and to interact with transfer RNA (tRNA) during translation by providing petidyltransferase activity (large subunit). Whereas the two rRNA subunits in eukaryotes have sedimentation coefficiency values of 40S (Svedberg units) and 60S, those in bacteria measure 30S and 50S, respectively. In virtually all organisms, the small rRNA subunit (40S in eukaryotes and 30S in bacteria) contains a single RNA species (i.e., 18S rRNA in eukaryotes and 16S rRNA in bacteria); the large rRNA subunit (60S) in eukaryotes comprises three RNA species (5S, 5.8S, and 25/28S rRNA), while that (50S) in bacteria contains two RNA species (5S and 23S rRNA). Although bacteria do not undergo meiosis or mitosis and do not require cellular fusion to initiate reproduction (as bacteria are not diploid), many bacteria do involve a cellto-cell transfer of genomic DNA by various mechanisms. These mechanisms may range from the uptake of exogenous DNA from their environment (a process called transformation) and the integration of a bacteriophage introduces

foreign DNA into the chromosome (a process called transduction), to the acquisition of DNA through direct cell contact (a process called conjugation). The incorporation of genes and DNA from other bacteria or the environment into the recipient cell’s DNA is also called horizontal gene transfer. While DNA transfer occurs less frequently per individual bacterium than that among eukaryotes involving obligate sexual reproduction, the much shorter generation times and high numbers associated with bacteria can make the DNA transfer a significant contributor to the evolution of bacterial populations. Gene transfer is vital to the development of antibiotic resistance in bacteria as it allows the rapid transfer of resistance genes between different pathogens. Regardless of genome size, most organisms show a mutation rate on the order of one mutation per genome per generation. Given their very short generation times ( two-cycle increase in the Cq for the extracted vaginal DNA sample compared to the no-template controls.133,134 Detection. After separation on 2% agarose gels, conventional PCR products are typically visualized by staining 5 µL of amplified product with ethidium bromide and placing the

gel under UV light.13,14,37,119–122,124 Specificity is confirmed by detection of the appropriate sized PCR product,124 which can be sequenced as an additional control.14,33,37 For detection in qPCR assays, the LightCycler (Roche),132 Stratagene MX 3000P,24,135 RotorGene 3000,130 and ABI platforms (Applied Biosystems) 39,94,118,133,134,136 have been used. In assays using SybrGreen I, analysis of the melting curve can be carried out after amplification to distinguish the targeted PCR product from aspecific nontargeted PCR products. For quantification, a standard curve is made. DNA, extracted from A. vaginae cultured on TSA + 5% sheep blood (Becton Dickinson) and diluted in a 1/10 series, can be used as standards.39,132 Alternatively, a serial diluted suspension of cloned 16S rRNA24,133,135 or cpn60118 plasmids can be used. 4.2.2.2.7  Sequence-Based Identification In cases were A. rimae, A. vaginae, or Atopobium sp. oral clone C019 isolates could not be identified or were misidentified using conventional techniques, the 16S rRNA gene was (partially) sequenced to reach identification.8,13,56–58

TABLE 4.1 Primers and Probes Used for Amplification and Detection of Atopobium spp. Primer and Probe Sequences (5′–3′)

Target Sequence

Target Species

Reference

GGTGAAGCAGTGGAAACACT ATTCGCTTCTGCTCGCGCA

16S rRNA gene

A. vaginae

39

CCCTATCCGCTCCTGATACC CCAAATATCTGCGCATTTCA VIC-GCAGGCTTGAGTCTGGTAGGGGA-TAMRA

16S rRNA gene

A. vaginae

24,135

GGTCGGTCTCTCAACCCGG TCATGGCCCAGAAGACCGCC FAM-CAGATTTAACTCCTGACCTAACAGACC-TAMRA

16S rRNA gene

A. vaginae

32,131

GCGAATATGGGAAAGCTCCG GAGCGGATAGGGGTTGAGC

16S rRNA gene

A. vaginae

120,122

TAGGTCAGGAGTTAAATCTG TCATGGCCCAGAAGACCGCC

16S rRNA gene

A. vaginae

123,124

Cpn60

A. vaginae

118

CAGCGTTCCTGTTACTCC GCCACTTGAAGGTCTTGC cgcgatcATTCCACGGTTGCACATAACATGCgatcgcga

16S rRNA gene

A. vaginae

130

GGTCGGTCTCTCAACCC CTCCTGACCTAACAGACC

16S rRNA gene

A. vaginae

14

GCAGGGACGAGGCCGCAA GTGTTTCCACTGCTTCACCTAA

16S rRNA gene

Atopobium spp.

37,121

TAGGCGGTYTGTTAGGTCAGGA CCTACCAGACTCAAGCCTGC FAM-CTCAACCCCTATCCGCTCCTGAT-TAMRA

16S rRNA gene

Atopobium spp.b

133,134

GGGTTGAGAGACCGACC CGGRGCTTCTTCTGCAGG

16S rRNA gene

Atopobium clusterc

94,132

CTTTGCAACCAACAATGACACC CAGCGCATACGGTAAGCGTAC

a b c

Uppercase letters indicate homology to the target sequence. Atopobium spp.: Atopobium sp. AY738657, AY738658. Atopobium cluster: Members of the genera Atopobium, Collinsella, Eggerthella, and Coriobacterium.

Atopobium

39

TABLE 4.2 Primers Used for Amplification of the 16S rRNA Gene Primer

Sequence 5′–3′

Amplicon Size

Reference

fD1 rP2

AGAGTTTGATCCTGGCTCAG ACGGCTACCTTGTTACGACTT

1454 bp

57

8FPL 806R

AGTTTGATCCTGGCTCAG GGACTACCAGGGTATCTAAT

798 bp

56

10f 534r

AGTTTGATCCTGGCTCAG ATTACCGCGGCTGCTGG

524 bp

13

515FPL 13B

TGCCAGCAGCCGCGGTAA AGGCCCGGGAACGTATTCAC

785 bp

56

Anaerobic blood cultures or swabs can be (sub)cultured on a rich blood agar (e.g., Columbia agar with 5% horse blood3) and incubated anaerobically for 2–5 days at 37°C (see growth requirements). For DNA extraction, commercial kits such as the QIAamp tissue kit (QIAGEN)57 or GenElute Bacterial Genomic DNA kit56 can be used, preferentially after pretreatment with mutanolysin. Alternatively, DNA is extracted from a colony by simple alkaline extraction.139 For sequencing purposes, 16S rRNA PCR amplification can be performed using 16S rRNA universal primers, as listed in Table 4.2. To obtain pure amplicons, excess primers and nucleotides are removed using a commercial kit (QIAquick PCR Purification Kit, Qiagen).13 Thereafter, the 16S rRNA gene amplicons are sequenced, either partially58 or fully.8,57,58 Commercial packages such as the MicroSeq® Full Gene or 500 16S rRNA Bacterial Identification Kit (PE Applied Biosystems) can be used.58 For identification, the sequence is compared to a public database (e.g., Genbank) or a curated database (e.g., SmartGene IDNS Bacteria) by using analysis software (BLAST version 2.2.957 or SmartGene [www.idnssmartgene.com]).

4.3  FUTURE PERSPECTIVES Highly sensitive high throughput analysis through deep pyrosequencing will further enlighten our knowledge of bacterial diversity and ecological dynamics associated with oral, reproductive tract, and intestinal tract infection(s). Knowledge obtained by this metagenomic approach might then be translated into rapid diagnostic assays for profiling a selected set of microbial communities in a lab-based setting. In case of bacterial vaginosis, distinct vaginal microflora patterns may be found in both intermediate and bacterial vaginosis microflora, thereby possibly differentiating intermediate microflora regressing to normal or developing to bacterial vaginosis and differentiating bacterial vaginosis microflora with different pathologic sequelae. Furthermore, molecular diagnosis of bacterial vaginosis, with a central role for A. vaginae, is now pending.37,118,133,135 Species-specific quantitative PCR techniques are being patented and will become commercially available as diagnostic assays. Also, ongoing sequence analysis of the whole genome

of A. vaginae within the A. vaginae Genome Project (http:// med.stanford.edu/sgtc/research/atopobium_vaginae.html) will probably reveal as yet undescribed virulence factors and will also improve our understanding of the pathogenic role of A. vaginae in reproductive tract infections. To explore the role of A. parvulum and A. rimae in oral health and disease, microarrays like the TNO oral microarray (O-chip, TNO, Zeist, The Netherlands) or the HOMIM (http://mim.forsyth.org/homim.html) offer the possibility to detect about 300 oral species. In cases of bacteremia, even if Atopobium spp. are included in future commercial biochemical tests, sequencing the 16S rRNA gene or the cpn60 gene will remain the golden standard to identify these species, as they are generally slow growing and biochemically nonreactive.12

REFERENCES











1. Collins, M.D., and Wallbanks, S., Comparative sequence analyses of the 16S rRNA genes of Lactobacillus minutus, Lactobacillus rimae and Streptococcus parvulus: Proposal for the creation of a new genus Atopobium, FEMS Microbiol. Lett., 74, 235, 1992. 2. Bailey, G.D., and Love, D.N., Eubacterium-Fossor sp. nov, an agar-corroding organism for normal pharynx and oral and respiratory-tract lesions of horses, Int. J. Syst. Bacteriol., 36, 383, 1986. 3. Rodriguez Jovita, M. et al., Characterization of a novel Atopobium isolate from the human vagina: Description of Atopobium vaginae sp. nov., Int. J. Syst. Bacteriol., 49, 1573, 1999. 4. Kageyama, A., Benno, Y., and Nakase, T., Phylogenic and phenotypic evidence for the transfer of Eubacterium fossor to the genus Atopobium as Atopobium fossor comb. nov., Microbiol. Immunol. 43, 389, 1999. 5. Rainey, F.A., Weiss, N., and Stackebrandt, E., Coriobacterium and Atopobium are phylogenetic neighbors within the Actinomycetes line of descent, Syst. Appl. Microbiol., 17, 202, 1994. 6. Stackebrandt, E., Rainey, F.A., and WardRainey, N.L., Propo­ sal for a new hierarchic classification system, Actinobacteria classis nov., Int. J. Syst. Bacteriol., 47, 479, 1997. 7. Olsen, I.E. et al., Lactobacillus uli sp. nov. and Lactobacillus rimae sp. nov. from the human gingival crevice and emended descriptions of Lactobacillus minutus and Streptococcus parvulus, Int. J. Syst. Bacteriol., 41, 261, 1991.

40













8. Geissdörfer, W. et al., Tuboovarian abscess caused by Atopobium vaginae following transvaginal oocyte recovery, J. Clin. Microbiol., 41, 2788, 2003. 9. Ferris, M.J. et al., Association of Atopobium vaginae, a recently described metronidazole resistant anaerobe, with bacterial vaginosis, BMC Infect. Dis., 4, 5, 2004. 10. Cato, E.P. et al., Transfer of Peptostreptococcus parvulus (Weinberg, Nativelle, and Prevot 1937) Smith 1957 to the gGenus Streptococcus as Streptococcus parvulus (Weinberg, Nativelle, and Prevot 1937) comb. nov., nom. rev., emend., Int. J. Syst. Bacteriol., 33, 82, 1983. 11. De Backer, E. et al., Antibiotic susceptibility of Atopobium vaginae, BMC Infect. Dis., 6, 51, 2006. 12. Downes, J. et al., Characterisation of Eubacterium-like strains isolated from oral infections, J. Med. Microbiol., 50, 947, 2001. 13. Verhelst, R. et al., Cloning of 16S rRNA genes amplified from normal and disturbed vaginal microflora suggests a strong association between Atopobium vaginae, Gardnerella vaginalis and bacterial vaginosis, BMC Microbiol., 4, 16, 2004. 14. Burton, J.P. et al., Detection of Atopobium vaginae in postmenopausal women by cultivation-independent methods warrants further investigation, J. Clin. Microbiol., 42, 1829, 2004. 15. Swidsinski, A. et al., An adherent Gardnerella vaginalis biofilm persists on the vaginal epithelium after standard therapy with oral metronidazole, Am. J. Obstet. Gynecol., 198, 97e1, 2008. 16. Sobel, J. D. et al., Bacterial vaginosis, Annu. Rev. Med., 51, 349, 2000. 17. Hillier, S.L. et al., The role of bacterial vaginosis and vaginal bacteria in amniotic fluid infection in women in preterm labor with intact fetal membranes, Clin. Infect. Dis., 20, S276, 1995. 18. Hillier, S.L. et al., Association between bacterial vaginosis and preterm delivery of a low-birth-weight infant. The Vaginal Infections and Prematurity Study Group, N. Engl. J. Med., 333, 1737, 1995. 19. Hillier, S.L. et al., Role of bacterial vaginosis-associated microorganisms in endometritis, Am. J. Obstet. Gynecol., 175, 435, 1996. 20. Hillier, S.L. et al., A case-control study of chorioamnionic infection and histologic chorioamnionitis in prematurity, N. Engl. J. Med., 319, 972, 1988. 21. Leitich, H. et al., Bacterial vaginosis as a risk factor for preterm delivery: A meta-analysis, Am. J. Obstet. Gynecol., 189, 139, 2003. 22. Sewankambo, N. et al., HIV-1 infection associated with abnormal vaginal flora morphology and bacterial vaginosis, Lancet, 350, 546, 1997. 23. Krohn, M.A. et al., The severity of fetal membrane infection and pregnancy complications, Ann. Epidemiol., 3, 78, 1993. 24. Menard, J.P. et al., High vaginal concentrations of Atopobium vaginae and Gardnerella vaginalis in women undergoing preterm labor, Obstet. Gynecol., 115, 134, 2010. 25. Wiesenfeld, H.C. et al., Bacterial vaginosis is a strong predictor of Neisseria gonorrhoeae and Chlamydia trachomatis infection, Clin. Infect. Dis., 36, 663, 2003. 26. Cherpes, T.L. et al., Association between acquisition of herpes simplex virus type 2 in women and bacterial vaginosis, Clin. Infect. Dis., 37, 319, 2003. 27. Atashili, J. et al., Bacterial vaginosis and HIV acquisition: A meta-analysis of published studies, AIDS, 22, 1493, 2008.

Molecular Detection of Human Bacterial Pathogens 28. Sha, B.E. et al., Female genital-tract HIV load correlates inversely with Lactobacillus species but positively with bacterial vaginosis and Mycoplasma hominis, J. Infect. Dis., 191, 25, 2005. 29. Cherpes, T.L. et al., Genital tract shedding of herpes simplex virus type 2 in women: Effects of hormonal contraception, bacterial vaginosis, and vaginal group B Streptococcus colonization, Clin. Infect. Dis., 40, 1422, 2005. 30. Josey, W.E., and Schwebke, J.R., The polymicrobial hypothesis of bacterial vaginosis causation: A reassessment, Int. J. STD AIDS, 19, 152, 2008. 31. Hay, P. Recurrent bacterial vaginosis, Curr. Opin. Infect. Dis., 22, 82, 2009. 32. Bradshaw, C.S. et al., The association of Atopobium vaginae and Gardnerella vaginalis with bacterial vaginosis and recurrence after oral metronidazole therapy, J. Infect. Dis., 194, 828, 2006. 33. Fredricks, D.N., Fiedler, T.L., and Marrazzo, J.M., Molecular identification of bacteria associated with bacterial vaginosis, N. Engl. J. Med., 353, 1899, 2005. 34. Fredricks, D.N., and Marrazzo, J.M., Molecular methodology in determining vaginal flora in health and disease: Its time has come, Curr. Infect. Dis. Rep., 7, 463, 2005. 35. Hillier, S.L. et al., The normal vaginal flora, H2O2-producing lactobacilli, and bacterial vaginosis in pregnant women, Clin. Infect. Dis., 16, S273, 1993. 36. Rosenstein, I.J. et al., Effect of intravaginal clindamycin cream on pregnancy outcome and on abnormal vaginal microbial flora of pregnant women, Infect. Dis. Obstet. Gynecol., 8, 158, 2000. 37. Fredricks, D.N., Targeted PCR for detection of vaginal bacteria associated with bacterial vaginosis, J. Clin. Microbiol., 45, 3270, 2007. 38. Thies, F.L., Konig, W., and Konig, B., Rapid characterization of the normal and disturbed vaginal microbiota by application of 16S rRNA gene terminal RFLP fingerprinting, J. Med. Microbiol., 56, 755, 2007. 39. De Backer, E. et al., Quantitative determination by real-time PCR of four vaginal Lactobacillus species, Gardnerella vaginalis and Atopobium vaginae indicates an inverse relationship between L. gasseri and L. iners, BMC Microbiol., 7, 115, 2007. 40. Verstraelen, H.M. et al., Culture-independent analysis of vaginal microflora: The unrecognized association of Atopobium vaginae with bacterial vaginosis, Am. J. Obstet. Gynecol., 191, 1130, 2004. 41. Gravett, M.G. et al., Preterm labor associated with subclinical amniotic fluid infection and with bacterial vaginosis, Obstet. Gynecol., 67, 229, 1986. 42. Watts, D.H. et al., The association of occult amniotic fluid infection with gestational age and neonatal outcome among women in preterm labor, Obstet. Gynecol., 79, 351, 1992. 43. Hebb, J.K. et al., Detection of novel organisms associated with salpingitis, by use of 16S rDNA polymerase chain reaction, J. Infect. Dis., 190, 2109, 2004. 44. Ferris, M.J. et al., Cultivation-independent analysis of changes in bacterial vaginosis flora following metronidazole treatment, J. Clin. Microbiol., 45, 1016, 2007. 45. Moreau-Marquis, S., Stanton, B.A., and O’Toole, G.A., Pseudomonas aeruginosa biofilm formation in the cystic fibrosis airway, Pulm. Pharmacol., Ther., 21, 595, 2008. 46. Prince, A.A. et al., Prevalence of biofilm-forming bacteria in chronic rhinosinusitis, Am. J. Rhinol., 22, 239, 2008.

Atopobium 47. Davies, D. et al., Understanding biofilm resistance to antibacterial agents, Nat. Rev. Drug Discov., 2, 114, 2003. 48. Swidsinski, A. et al., Adherent biofilms in bacterial vaginosis, Obstet. Gynecol., 106, 1013, 2005. 49. Costerton, J.W., Stewart, P.S., and Greenberg, E.P., Bacterial biofilms: A common cause of persistent infections, Science, 284, 1318, 1999. 50. Hale, L.P., Swidsinski, A., and Mendling, W., Bacteria associated with bacterial vaginosis, N. Engl. J. Med., 354, 202, 2006. 51. Libby, E.K. et al., Atopobium vaginae triggers an innate immune response in an in vitro model of bacterial vaginosis, Microbes Infect., 10, 439, 2008. 52. Barrett, S., and Taylor, C., A review on pelvic inflammatory disease, Int. J. STD AIDS 16, 715; quiz 721, 2005. 53. Bebear, C., and de Barbeyrac, B., Genital Chlamydia trachomatis infections, Clin. Microbiol. Infect., 15, 4, 2009. 54. Reyes, P.A. et al., Apoptosis related genes expressed in cultured Fallopian tube epithelial cells infected in vitro with Neisseria gonorrhoeae, Biol. Res., 40, 319, 2007. 55. Soper, D.E. et al., Pelvic inflammatory disease, Infect. Dis. Clin. North Am., 8, 821, 1994. 55a. Yamagishi, Y. et al., A case of uterine endometritis caused by Atopobium vaginae, J. Infect. Chemother., 2010 Aug 14 [Epub ahead of print]. 56. Chung, H.Y. et al., A case of bacteremia by Atopobium rimae in a patient with liver cirrhosis, Korean J. Lab. Med., 27, 351, 2007. 57. Angelakis, E.M. et al., Human case of Atopobium rimae bacteremia, Emerg. Infect. Dis., 15, 354, 2009. 58. Salimnia, H. et al., Sepsis associated with a new Atopobium species, provisionally named Atopobium detroiti: Case report and review of the current status of the species Atopobium, Scand. J. Infect. Dis., 40, 679, 2008. 59. Dewhirst, F.E. et al., Characterization of novel human oral isolates and cloned 16S rDNA sequences that fall in the family Coriobacteriaceae: Description of Olsenella gen. nov., reclassification of Lactobacillus uli as Olsenella uli comb. nov. and description of Olsenella profusa sp. nov., Int. J. Syst. Evol. Microbiol., 51, 1797, 2001. 60. Hughes, F.J., and McNab, R., Oral malodour—a review, Arch. Oral Biol., 53, S1, 2008. 61. Kazor, C.E. et al., Diversity of bacterial populations on the tongue dorsa of patients with halitosis and healthy patients, J. Clin. Microbiol., 41, 558, 2003. 62. Persson, S.J. et al., The formation of hydrogen sulfide and methyl mercaptan by oral bacteria, Oral Microbiol. Immunol., 5, 195, 1990. 63. Riggio, M.P. et al., Molecular identification of bacteria on the tongue dorsum of subjects with and without halitosis, Oral Dis., 14, 251, 2008. 64. Haraszthy, V.I. et al., Identification of oral bacterial species associated with halitosis, J. Am. Dent. Assoc., 138, 1113, 2007. 65. Wolff, M.S., and Larson, C., The cariogenic dental biofilm: Good, bad or just something to control? Braz. Oral Res., 23, S31, 2009. 66. Loesche, W.J. et al., The specific plaque hypothesis and the antimicrobial treatment of periodontal disease, Dent. Update, 19, 70, 1992. 67. Theilade, E. et al., The non-specific theory in microbial etiology of inflammatory periodontal diseases, J. Clin. Periodontol., 13, 905, 1986. 68. Marsh, P.D. et al., Microbial ecology of dental plaque and its significance in health and disease, Adv. Dent. Res., 8, 263, 1994.

41 69. Aas, J.A. et al., Bacteria of dental caries in primary and permanent teeth in children and young adults, J. Clin. Microbiol., 46, 1407, 2008. 70. Moore, W.E. et al., Bacteriology of experimental gingivitis in young adult humans, Infect. Immun., 38, 651, 1982. 71. Moore, W.E. et al., Bacteriology of experimental gingivitis in children, Infect. Immun., 46, 1, 1984. 72. Moore, L.V. et al., Bacteriology of human gingivitis, J. Dent. Res., 66, 989, 1987. 73. Socransky, S.S., and Haffajee, A.D., Microbial mechanisms in the pathogenesis of destructive periodontal diseases: A critical assessment, J. Periodont. Res., 26, 195, 1991. 73a. Svensäter, G. et al., Biofilms in endodontic infections, Endod. Topics, 9, 27, 2004. 74. Ørstavik, D., and Pitt Ford, T.R., Essential Endodontology: Prevention and Treatment of Apical Periodontitis, 2nd edn, Blackwell Munksgaard, 2008. 75. Moore, W.E. et al., Bacteriology of severe periodontitis in young adult humans, Infect. Immun., 38, 1137, 1982. 76. Moore, W.E. et al., Comparative bacteriology of juvenile periodontitis, Infect. Immun., 48, 507, 1985. 77. Kumar, P.S. et al., Identification of candidate periodontal pathogens and beneficial species by quantitative 16S clonal analysis, J. Clin. Microbiol., 43, 3944, 2005. 78. Subramanian, K., and Mickel, A.K., Molecular analysis of persistent periradicular lesions and root ends reveals a diverse microbial profile, J. Endod., 35, 950, 2009. 79. Munson, M.A. et al., Molecular and cultural analysis of the microflora associated with endodontic infections, J. Dent. Res., 81, 761, 2002. 80. Siqueira, J.F. et al., Novel bacterial phylotypes in endodontic infections, J. Dent. Res., 84, 565, 2005. 81. Siqueira, J.F. et al., Bacteria in the apical root canal of teeth with primary apical periodontitis, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 107, 721, 2009. 82. Kato, N., and Kato, H., Molecular detection and identification of anaerobic bacteria, J. Infect. Chemother., 3, 5, 1997. 83. Olsen, G.J., and Woese, C.R., Ribosomal RNA: A key to phylogeny, FASEB J., 7, 113, 1993. 84. de Boer, R. et al., Improved detection of microbial DNA after bead-beating before DNA isolation, J. Microbiol. Methods, 80, 209, 2010. 85. Fliss, I. et al., A rapid and efficient method of lysis of Listeria and other gram-positive bacteria using mutanolysin, Biotechniques, 11, 456, 1991. 86. Moter, A., and Gobel, U.B., Fluorescence in situ hybridization (FISH) for direct visualization of microorganisms, J. Microbiol. Methods, 41, 85, 2000. 87. Glockner, F.O., Fuchs, B.M., and Amann, R., Bacterioplankton compositions of lakes and oceans: A first comparison based on fluorescence in situ hybridization, Appl. Environ. Microbiol., 65, 3721, 1999. 88. 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. 89. Amann, R., Fuchs, B.M., and Behrens, S., The identification of microorganisms by fluorescence in situ hybridisation, Curr. Opin. Biotechnol., 12, 231, 2001. 90. Jain, K.K. et al., Current status of fluorescent in-situ hybridisation, Med. Device Technol., 15, 14, 2004. 91. Levsky, J.M., and Singer, R.H., Fluorescence in situ hybridization: Past, present and future, J. Cell Sci., 116, 2833, 2003.

42 92. Harmsen, H.J. et al., Development of 16S rRNA-based probes for the Coriobacterium group and the Atopobium cluster and their application for enumeration of Coriobacteriaceae in human feces from volunteers of different age groups, Appl. Environ. Microbiol., 66, 4523, 2000. 93. Rigottier-Gois, L. et al., Fluorescent hybridisation combined with flow cytometry and hybridisation of total RNA to analyse the composition of microbial communities in human faeces using 16S rRNA probes, FEMS Microbiol. Ecol., 43, 237, 2003. 94. Matsuki, T. et al., Use of 16S rRNA gene-targeted groupspecific primers for real-time PCR analysis of predominant bacteria in human feces, Appl. Environ. Microbiol., 70, 7220, 2004. 95. Mah, K.W. et al., Effect of a milk formula containing probiotics on the fecal microbiota of asian infants at risk of atopic diseases, Pediatr. Res., 62, 674, 2007. 96. Kleessen, B. et al., Microbial and immunological responses relative to high-altitude exposure in mountaineers, Med. Sci. Sports Exerc., 37, 1313, 2005. 97. Socransky, S.S. et al., “Checkerboard” DNA–DNA hybridization, Biotechniques, 17, 788, 1994. 98. Nascimento, C. et al., DNA checkerboard method for bacterial pathogen identification in oral diseases, Int. J. Morphol., 24, 619, 2006. 99. Ximenez-Fyvie, L.A., Haffajee, A.D., and Socransky, S.S., Microbial composition of supra- and subgingival plaque in subjects with adult periodontitis, J. Clin. Periodontol., 27, 722, 2000. 100. Siqueira, J.F. et al., Endodontic infections: Concepts, paradigms, and perspectives, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 94, 281, 2002. 101. Socransky, S.S. et al., Microbial complexes in subgingival plaque, J. Clin. Periodontol., 25, 134, 1998. 102. Sakamoto, M., Umeda, M., and Benno, Y., Molecular analysis of human oral microbiota, J. Periodont. Res., 40, 277, 2005. 103. Colombo, A.P. et al., Clinical and microbiological features of refractory periodontitis subjects, J. Clin. Periodontol., 25, 169, 1998. 104. Haffajee, A.D. et al., Subgingival microbiota in healthy, well-maintained elder and periodontitis subjects, J. Clin. Periodontol., 25, 346, 1998. 105. Nikolaitchouk, N. et al., The lower genital tract microbiota in relation to cytokine-, SLPI- and endotoxin levels: Application of checkerboard DNA–DNA hybridization (CDH), APMIS, 116, 263, 2008. 106. Brito, L.C. et al., Use of multiple-displacement amplification and checkerboard DNA–DNA hybridization to examine the microbiota of endodontic infections, J. Clin. Microbiol., 45, 3039, 2007. 107. Smith, G.L., Socransky, S.S., and Smith, C.M., Rapid method for the purification of DNA from subgingival microorganisms, Oral Microbiol. Immunol., 4, 47, 1989. 108. Paster, B.J., Bartoszyk, I.M., and Dewhirst, F., Identification of oral streptococci using PCR-based, reverse-capture, checkerboard hybridization, Methods Cell Sci., 20, 223, 1998. 109. Paster, B.J. et al., Bacterial diversity in necrotizing ulcerative periodontitis in HIV-positive subjects, Ann. Periodontol., 7, 8, 2002. 110. Siqueira, J.F., Jr., and Rocas, I.N., The microbiota of acute apical abscesses, J. Dent. Res., 88, 61, 2009. 111. Schena, M. et al., Parallel human genome analysis: Microarraybased expression monitoring of 1000 genes, Proc. Natl. Acad. Sci. USA, 93, 10614, 1996.

Molecular Detection of Human Bacterial Pathogens 112. Mothershed, E.A., and Whitney, A.M., Nucleic acid-based methods for the detection of bacterial pathogens: Present and future considerations for the clinical laboratory, Clin. Chim. Acta 363, 206, 2006. 113. Palmer, C. et al., Rapid quantitative profiling of complex microbial populations, Nucleic Acids Res., 34, 2006. 114. Olsen, I. et al., Cultivated and not-yet-cultivated bacteria in oral biofilms, Microb. Ecol. Health Dis., 65, 2009. 115. Preza, D. et al., Microarray analysis of the microflora of root caries in elderly, Eur. J. Clin. Microbiol. Infect. Dis., 28, 509, 2009. 116. Preza, D. et al., Diversity and site-specificity of the oral microflora in the elderly, Eur. J. Clin. Microbiol. Infect. Dis., 28, 1033, 2009. 117. Colombo, A.P. et al., Comparisons of subgingival microbial profiles of refractory periodontitis, severe periodontitis, and periodontal health using the human oral microbe identification microarray, J. Periodontol., 80, 1421, 2009. 118. Dumonceaux, T.J. et al., Multiplex detection of bacteria associated with normal microbiota and with bacterial vaginosis in vaginal swabs by use of oligonucleotide-coupled fluorescent microspheres, J. Clin. Microbiol., 47, 4067, 2009. 119. Burton, J.P. et al., A preliminary survey of Atopobium vaginae in women attending the Dunedin gynaecology out-patients clinic: Is the contribution of the hard-to-culture microbiota overlooked in gynaecological disorders? Aust. N. Z. J. Obstet. Gynaecol., 45, 450, 2005. 120. Weyers, S. et al., Microflora of the penile skin-lined neovagina of transsexual women, BMC Microbiol., 9, 102, 2009. 121. Marrazzo, J.M. et al., Relationship of specific vaginal bacteria and bacterial vaginosis treatment failure in women who have sex with women, Ann. Intern. Med., 149, 20, 2008. 122. Verstraelen, H. et al., Gene polymorphisms of Toll-like and related recognition receptors in relation to the vaginal carriage of Gardnerella vaginalis and Atopobium vaginae, J. Reprod. Immunol., 79, 163, 2009. 123. Ferris, M.J., Masztal, A., and Martin, D.H., Use of species-directed 16S rRNA gene PCR primers for detection of Atopobium vaginae in patients with bacterial vaginosis, J. Clin. Microbiol., 42, 5892, 2004. 124. Haggerty, C.L. et al., Clinical characteristics of bacterial vaginosis among women testing positive for fastidious bacteria, Sex. Transm. Infect., 85, 242, 2009. 125. Higuchi, R. et al., Kinetic PCR analysis: Real-time monitoring of DNA amplification reactions, Biotechnology (N Y), 11, 1026, 1993. 126. Arya, M. et al., Basic principles of real-time quantitative PCR, Expert. Rev. Mol. Diagn., 5, 209, 2005. 127. Koch, W.H. et al., Technology platforms for pharmacogenomic diagnostic assays, Nat. Rev. Drug. Discov., 3, 749, 2004. 128. Kim, Y., Sohn, D., and Tan, W., Molecular beacons in biomedical detection and clinical diagnosis, Int. J. Clin. Exp. Pathol., 1, 105, 2008. 129. Abravaya, K. et al., Molecular beacons as diagnostic tools: Technology and applications, Clin. Chem. Lab. Med., 41, 468, 2003. 130. Trama, J.P. et al., Rapid detection of Atopobium vaginae and association with organisms implicated in bacterial vaginosis, Mol. Cell. Probes, 22, 96, 2008. 131. Tabrizi, S.N. et al., Prevalence of Gardnerella vaginalis and Atopobium vaginae in virginal women, Sex. Transm. Dis., 33, 663, 2006.

Atopobium 132. Biagi, E. et al., Quantitative variations in the vaginal bacterial population associated with asymptomatic infections: A realtime polymerase chain reaction study, Eur. J. Clin. Microbiol. Infect. Dis., 28, 281, 2009. 133. Fredricks, D.N. et al., Changes in vaginal bacterial concentrations with intravaginal metronidazole therapy for bacterial vaginosis as assessed by quantitative PCR, J. Clin. Microbiol., 47, 721, 2009. 134. Mitchell, C.M et al., Comparison of oral and vaginal metronidazole for treatment of bacterial vaginosis in pregnancy: Impact on fastidious bacteria, BMC Infect. Dis., 9, 89, 2009. 135. Menard, J.P. et al., Molecular quantification of Gardnerella vaginalis and Atopobium vaginae loads to predict bacterial vaginosis, Clin. Infect. Dis., 47, 33, 2008.

43 136. Matsuda, K. et al., Establishment of an analytical system for the human fecal microbiota, based on reverse transcriptionquantitative PCR targeting of multicopy rRNA molecules, Appl. Environ. Microbiol., 75, 1961, 2009. 137. Yamamoto, T. et al., Bacterial populations in the vaginas of healthy adolescent women, J. Pediatr. Adolesc. Gynecol., 22, 11, 2009. 138. Zhou, X. et al., Characterization of vaginal microbial communities in adult healthy women using cultivation-independent methods, Microbiol. Sgm., 150, 2565, 2004. 139. Vaneechoutte, M. et al., Isolation of Moraxella canis from an ulcerated metastatic lymph node, J. Clin. Microbiol., 38, 3870, 2000.

5 Bifidobacterium Abelardo Margolles, Patricia Ruas-Madiedo, Clara G. de los Reyes-Gavilán, Borja Sánchez, and Miguel Gueimonde CONTENTS 5.1 Introduction........................................................................................................................................................................ 45 5.1.1 Classification and Taxonomy.................................................................................................................................. 45 5.1.2 Biology and Physiology.......................................................................................................................................... 46 5.1.3 Clinical Features and Pathogenesis........................................................................................................................ 49 5.1.4 Diagnosis................................................................................................................................................................ 51 5.1.4.1 Phenotypic Techniques............................................................................................................................ 51 5.1.4.2 Molecular Techniques.............................................................................................................................. 52 5.2 Methods.............................................................................................................................................................................. 53 5.2.1 Sample Preparation................................................................................................................................................. 53 5.2.2 Detection Procedures.............................................................................................................................................. 53 5.3 Conclusion and Future Perspectives................................................................................................................................... 54 References.................................................................................................................................................................................... 54

5.1  INTRODUCTION 5.1.1  Classification and Taxonomy The genus Bifidobacterium includes high G + C gram-­positive nonspore-forming, nonmotile, and nonfilamentous polymorphic rod-shaped bacteria that can display slight bends or a large variety of branchings, the slightly bifurcated club-shaped or spatulated extremities being the most commonly found. They can be organized singly or in chains, in star-like aggregates, in “V,” or in palisade arrangements when grown in vitro. They are strictly anaerobic, although some species, such as Bifidobacterium animalis and Bifidobacterium psychraerophilum, can tolerate moderately high oxygen concentrations, and they have a fermentative metabolism.1,2 Bifidobacteria were first described at the beginning of the twentieth century and were included among the family Lactobacillaceae.3 In 1924, Lactobacillus bifidum was reclassified as the new genus Bifidobacterium by Orla-Jensen.1 Currently, over 30 species are included in the genus Bifidobacterium. They form a close phylogenetic group and show high similarity (>93%) in the 16S rRNA sequences.4 The Bifidobacterium genus is included in the phylum Actino­ bacteria, class Actinobacteria, subclass Actinobacteridae, order Bifidobacteriales, and family Bifido­bacteriaceae. Phylogenetic studies, based on sequence comparison of 16S rRNA sequences and housekeeping genes, have clustered some Bifidobacterium species in six phylogenetic clusters named Bifidobacterium boum, Bifidobacterium

asteroides, Bifidobacterium adolescentis, Bifidobacterium longum, Bifidobacterium pullorum, and Bifidobacterium pseudolongum.5 Other species have displayed a large phylogenetic distance and have not been assigned to any specific cluster (Figure 5.1). The species included in the genus Bifidobacterium are B. adolescentis, B. angulatum, B. animalis, B.  asteroides, B.  bifidum, B.  bombi, B.  boum, B.  breve,  B. catenulatum, B.  choerinum, B. coryneforme, B. cuniculi, B. dentium, B. gallicum, B. gallinarum, B. indicum, B.  longum, B.  magnum, B.  merycicum, B.  minimum, B.  mongoliense, B.  pseudocatenulatum, B.  pseudolongum, B. psychraerophilum, B. pullorum, B. ruminantium, B. saeculare, B. scardovii, B. subtile, B. thermacidophilum, B. thermophilum, and B. tsurumiense.6–9 Further­more, B. animalis comprises of two subspecies (animalis and lactis), as well as B. pseudolongum (subsp. globosum and pseudolongum), and B. thermacidophilum (subsp. thermoacidophilum and porcinum), whereas the species B.  longum is subdivided in the subspecies longum, infantis, and suis (Figure 5.1). Bifidobacterium species can be isolated from a limited number of habitats—that is, human and animal gastrointestinal tract (GIT), food, insect intestine, and sewage.10,11 Among the species most commonly found in human intestine and feces are B. catenulatum, B. pseudocatenulatum, B. adolescentis, B.  longum, B. pseudolongum, B. breve, B. angulatum, B. bifidum, and B. dentium, whereas B. animalis is the species more often found in functional foods.12 In this respect, it is of key importance to remark that a clear 45

46

e AB433856 B. mongoliens

D86183 B. dentium

ga llin ar um

D8 61 9 B . pu llorum 1 D8619 6

B.

4108 AY17 ilum eroph ychra B . ps

B.

B. adolescentis group

1 874 M5 um inim B. m 8 932 78 D8 93 are D8 cul tile sae B. sub

B.

B. pullorum group

B. adolescentis NC008618 B . ru min anti me um D B. ryc 8619 icu an 7 mD gu 861 la 92 tu m D8 61 82

Molecular Detection of Human Bacterial Pathogens

tum ula 7 ten a m D8618 c . tenulatu

BB . pseudoca

B . scard ovii AJ3 07005

24 S836 ifdum b . B

B. longum group

B. boum group

U10151 ophilum B . therm

B. breve AB006658

s . infanti m subsp 86184 B . longu D s i u s sp. 3 sub 874 gum 9 um M5 lon 873 g n . 5 lo p . s B b M su m gu n lo B.

B. thermacidophilum subsp. thermacidop hilum

AY 14 84 70

B. asteroides group

58734 iculi M B . cun

B. magnum M58740

B . gall icum D 86189

88 61 D8 m cu 30 di 587 in sM B. 49 ide ero 1275 ast i EU B. omb B. b 4110 se AB2 umien B . tsur

B . B . bo um D AB016246 the rm 8619 aci 0 do ph ilu m B. c sub oryn sp. efor po me rci M5 nu 873 m 3

5 18 86 .D bsp s su 3 ali X8951 im . lactis an s subsp B . B . animali

B . p seud B. olon pse gum ud subs B o p . l g o . lo ngu bosu ch mD m oe 8619 sub rin 4 sp u . p m seu do lon g u m D8 61 95

D8 61 86

0.005

082 432 AF

B. pseudolongum group FIGURE 5.1  Evolutionary relationships of bifidobacteria.

ecological specialization seems to exist for different bifidobacterial taxa, and species distribution depends on host age and localization in the host (i.e., intestinal mucosa versus luminal/fecal environment).13,14 Furthermore, novel metagenomic studies have analyzed the bifidobacterial microbiota in the human gut and revealed that its biodiversity was much broader than had been previously expected,14,15 showing the existence of uncharacterized phylotypes that have passed unnoticed until recently, and highlighting the importance of novel culture-independent approaches to complement the more traditional culture-dependent techniques.

5.1.2  Biology and Physiology As stated above, the natural habitat of the genus Bifidobac­ terium is the GIT of mammals, birds, and insects. The human GIT hosts a very complex microbiota that plays an important role on the health and wellbeing of the host.16 This microbial ­community reaches one of the highest cell densities recorded for any ecosystem, although its taxonomical diversity is one of the lowest. A small number of taxa dominate this ecosystem in adult humans, mainly representatives of Bacteroidetes and Firmicutes divisions.17 The phylum Actinobacteria, including the Bifidobacteriaceae family, is also present.

Bifidobacterium

The GIT colonization of newborn babies starts at birth and increases in number and diversity in few days, bifidobacteria being one of the first and most abundant colonizers. Several factors affect this process, such as the type of delivery, the initial way of feeding (breast or infant formula), or the geographical region, among others. The number of bifidobacteria declines with the age of the host, remaining more or less stable during the adult life and tending to decrease in the elderly.18 In addition, intersubject species variability in bifidobacterial populations has been noticed, but low diversity is found between different GIT regions of the same subject, denoting scarce intrasubject variability.14 At the beginning of the twentieth century, Tissier3 described for the first time bifidobacterial strains (originally named Bacillus bifidus) isolated from breastfed infant feces. During recent years, an increasing number of articles have been published reporting on the bifidobacterial diversity of the GIT ecosystem. Commonly, these studies are based on fecal sampling, leading to the isolation of new strains. However, in the fecal material, transitory bacteria derived from the diet are usually present, and therefore not representing the mucosaadhered microbiota.19 For this reason, different GIT sections, such as colonic mucosa obtained from biopsies of healthy individuals, are also used to explore the bifidobacterial diversity of the human microbiota.15 In addition, Bifidobacterium strains can also be isolated from human saliva20 and breast milk,21 as well as from sewage, in which bifidobacteria are used as indicators of fecal contamination.22 Regarding the physiology of this genus, the optimal growth temperature of most human isolated strains is about 36°C– 38°C, being higher for that of animal origin (41°C–43°C). Viability of bifidobacteria at acidic pH values is very variable but, in general, it can be considered that resistance of bifidobacteria to acidic pH is weak, with the exception of B. animalis.23 This species satisfactorily tolerates acidic pH and, in addition, the presence of oxygen, for these reasons being the most used Bifidobacterium spp. in fermented probiotic products. It has been suggested that the acid tolerance in Bifidobacterium is linked to the activity of the membrane-bound F0F1-ATPase, an enzyme responsible for the maintenance of the intracellular pH homeostasis in anaerobic bacteria through the ATPdependent extrusion of protons.24 The high F0F1-ATPase activity induction observed for this species appears to be responsible for its elevated acid resistance, this activity not being induced in nonresistant strains.25 Tolerance to bile salts is another crucial property for the persistence of bifidobacteria in the human GIT. Usually, deconjugated bile salts are more toxic than their corresponding taurine or glycine conjugates. The enzyme responsible for bile salts deconjugation is the bile salt hydrolase (BSH), but to date there is no solid evidence of a relationship between this enzyme and the levels of resistance to bile in bifidobacteria.26 Bifidobacteria have a specific pathway for the metabolism of sugars, the fructose-6-phosphate phosphoketolase (F6PPK) pathway or “bifido shunt” (Figure 5.2; Table 5.1), described for the first time in the work of Scardovi and Trovatelli.27 The key enzyme of this pathway is a xylulose-

47

5-phosphate/fructose-6-phosphate phosphoketolase (Xfp), which catalyzes the phosphorolytic cleavage of fructose-6phosphate and/or xylulose-5-phosphate. To date, two types of Xfp activities have been described in prokaryotes and eukaryotes—fructose-6-phosphate phosphoketolase (F6PPK, EC 4.1.2.22), which catalyzes the conversion of fructose-6phosphate to erythrose-4-phosphate and acetyl-phosphate; and xylulose-5-phosphate phosphoketolase (EC 4.1.2.9), which cleaves xylulose-5-phosphate, rendering acetyl-phosphate and glyceraldehyde-3-phosphate.28 In bifidobacteria, in addition to the F6PPK, a dual xylulose-5-phosphate/fructose6-phosphate phosphoketolase able to act over both fructose6-phosphate and xylulose-5-phosphate has been described.29 Bifidobacteria also present a transaldolase (Tal, EC 2.2.1.2) and a transketolase (Tkt, EC 2.2.1.1) able to convert the fructose-6-phosphate in acetyl-phosphate and glyceraldehyde-3phosphate. An acetate kinase (AckA, EC 2.7.2.1) allows the conversion of acetyl-phosphate into acetate with the concomitant production of ATP. The glyceraldehyde-3-phosphate, through the last reactions of the Embden–Meyerhof–Parnas (EMP) pathway, is metabolized to pyruvate and the activity of lactate dehydrogenase NAD-dependent (Ldh2, EC 1.1.1.27) converts this intermediate metabolite in lactate. In addition, pyruvate in combination with Co-A is converted into formate and acetyl-CoA by the activity of the formate C-acetyltransferase (Pfl, EC 2.3.1.54). Finally, acetyl-CoA is metabolized to ethanol, by means of the bifunctional acetaldehyde-CoA/alcohol dehydrogenase (Adh2, EC 1.1.1.1), and/ or to acetate by the activity of the phosphate acetyltransferase (Pta, EC 2.3.1.8). The theoretical ratio of the bifido shunt gives 1 mol of lactic acid and 1.5 moles of acetic acid per mol of glucose consumed, producing 2.5 moles of ATP without CO2 formation, although the production of formic acid and ethanol can modify this fermentation balance. The energetic balance of the bifido shunt (2.5 moles ATP) is higher than that obtained by lactic acid bacteria after the hexose metabolism through the EMP (or homofermentative pathway, 2  moles ATP) or through the pentose-phosphoketolase (or heterofermentative pathway, 1 mol ATP). The availability of several genome sequences of Bifidobacterium species provides genetic evidence to indicate that it is a prototrophic genus for some growth factors—that is, it is able to synthesize some vitamins (B complex, folic acid, thiamine, and nicotinate), aminoacids, and nucleotides. Genomic information also indicates that several species harbor genes required for metabolizing many monosaccharides or disaccharides through the fructose-6-phosphate phosphoketolase pathway. Analysis of the first bifidobacterial genome available, B. longum NCC2705,30 indicated that this strain possesses all the enzyme-coding genes necessary to metabolize several plant- and milk-derived oligosaccharides, containing fructose, galactose, N-acetyl-glucosamine, N-acetyl-galactosamine, arabinose, xylose, ribose, sucrose, lactose, cellobiose, and melibiose through this shunt. In fact, nearly 10% of the genes contained in the genome of this strain encode for proteins that are predicted to be involved in the carbohydrate metabolism and transport. This supposes,

48

Molecular Detection of Human Bacterial Pathogens

Glucose ATP

GlkA Glucose-6-P Gpi Fructose-6-P

Acetyl-P

Xfp Fructose-6-P

Erythrose-1-P

Sedoheptulose-7-P

ATP

Glyceraldehyde-3-P Tkt

Ribose-5-P

AckA

Tal

Xylulose-5-P

Xfp Acetate

Glyceraldehyde-3-P NADH + H+

Gap 1.3-Biphosphoglycerate Pgk

ATP

3-Phosphoglycerate

AckA

Gpm

ATP

2-Phosphoglycerate Eno

Acetyl-P

Phosphoenolpyruvate ATP

Pyk Pyruvate

Pfl

Ldh2

NAD*

Lactate

Pta Acetyl-CoA Adh2

Formate

Acetaldehyde Adh2

EMP pathway

NAD*

NAD*

Ethanol

FIGURE 5.2  Fructose-6-phosphate phosphoketolase pathway “Bifid shunt.”

theoretically, a selective advantage to colonize the colon, an environment poor in mono- and disaccharides because these are consumed by the host and microbiota in the upper GIT. Based on amino acid sequence identity, the presence of several glycolytic activities, including xylanases, arabinosidades, α-galactosidases, neopullanase, isomaltase, maltase, inulinase, β-galactosidases, β-glucosidases, hexosaminidases, and α-manosidases, as well as eight high-affinity oligosaccharide transporters, has been predicted.30 Similar gene arrangements have been seen after the genome annotation and comparison of the bifidobacterial species B. adolescentis ATCC15703, B. animalis subsp. lactis AD011, and B. longum subsp. infantis ATCC15697.31,32 However, comparative genome analysis revealed some differences, such as the presence of a fos gene cluster in B. animalis subsp. lactis AD011, very similar to the one described in B. breve UCC2003.33 This cluster is likely to be involved in fructooligosaccharide (FOS) catabolism, which is present in a panoply of food ingredients and supplements. FOS are composed of 4–60 fructose monomers linked by β(2–1) bonds, with a terminal glucose also connected by β(2–1)

linkage. This β(2–1) bond is not recognized as a substrate by intestinal enzymes, and thus FOS are able to reach the colon, where they stimulate the growth of beneficial bacteria, including bifidobacteria.34 This capacity of beneficially modulating the composition of the intestinal microbiota allows FOS to be considered as prebiotics.35 The presence of fos cluster reflects the metabolic adaptation of bifidobacteria to the human GIT conditions. Interestingly, comparative genomics revealed the presence of a specific genes cluster involved in the transport and metabolism of human milk oligosaccharides (HMO) in the strain B. longum subsp. infantis ATCC15697.31 This strain lacked certain enzymatic activities related to the metabolism of plant-derived oligosaccharides, such as arabinose and xylose, which are abundant in adult diets. This fact, together with the evidence of the presence of similar HMO-related gene clusters in infant gut metagenomes, suggests a specific adaptation of B. longum subsp. infantis ATCC15697 to the infant GIT.36 Thus, bifidobacteria are able to ferment a wide variety of carbohydrates. In addition, they are capable of

49

Bifidobacterium

TABLE 5.1 Accession Numbers of the Major Proteins Acting in Bifid Shunt in Some Available Bifidobacterium Genomes

GlkA Gpi Tal Tkt Xfp Gap Pgk Gpm Eno Pyk Ldh2 Pfl Pta AckA Adh2

B. adolescentis ATCC 15703

B. animalis subsp. lactis AD011

B. bifidum NCIMB 41171

B. longum subsp. longum NCC2705

B. longum subsp. infantis ATCC 15697

YP_909820 YP_909094 YP_909693 YP_909692 YP_909550 YP_909942 YP_909698 YP_909241 YP_909508 YP_909541 YP_909980 YP_909855 YP_909551 YP_909552 YP_909182

YP_002470082 YP_002469157 YP_002470219 YP_002470218 YP_002470341 YP_002469609 YP_002470227 YP_002469301 YP_002469894 YP_002470352 YP_002469660 YP_002469833 YP_002470342 YP_002470343 YP_002469236

ZP_03647167 ZP_03645763 ZP_03646448 ZP_03646447 ZP_03646196 ZP_03645995 ZP_03646458 ZP_03647102 ZP_03646131 ZP_03646166 ZP_03646455 ZP_03646658 ZP_03646187 ZP_03646186 ZP_03647027

NP_696841 NP_695484 NP_695898 NP_695899 NP_696135 NP_696527 NP_695890 NP_696806 NP_696193 NP_696160 NP_696472 NP_696127 NP_696142 NP_696143 NP_696730

YP_002322055 YP_002321916 YP_002322563 YP_002322564 YP_002323176 YP_002322378 YP_002322555 YP_002323591 YP_002323289 YP_002323198 YP_002322558 YP_002323169 YP_002323183 YP_002323184 YP_002323678

Additional information can be obtained through their corresponding accession number at http://www.ncbi.nlm.nih.gov/protein. GlkA, glucokinase; Gpi, glucose-6-phosphate isomerase; Tal, transaldolase; Tkt, transketolase; Xfp, xylulose-5-phosphate/fructose-6-phosphate phosphoketolase; Gap, glyceraldehyde-3-phosphate dehydrogenase C; Pgk, phosphoglycerate kinase; Gpm, phosphoglycerate mutase; Eno, enolase; Pyk, pyruvate kinase; Ldh2, lactate dehydrogenase; Pfl, formate acetyltransferase; Pta, Phosphate acetyltransferase; AckA, acetate kinase; Adh2, bifunctional acetaldehyde-CoA/alcohol dehydrogenase.

processing indigestible complex carbohydrates from our diet and from host origin.6,30,37–39 Hence, the availability of genome sequences of intestinal Bifidobacterium spp., together with the use of physiological experiments, has provided considerable insight into the adaptation of this genus to the human gut environment, ­especially regarding the utilization of dietary carbohydrates, resistance to bile and acid conditions, and interaction with the host.

5.1.3  Clinical Features and Pathogenesis It has been shown that Bifidobacterium is present at high levels in the gut microbiota mainly in infants but also in adults.40 Reduced levels of these microorganisms in the intestine of subjects suffering from different diseases, including colon cancer, irritable bowel syndrome, inflammatory bowel disease, atopic disease, or celiac disease, have been observed.41–45 For these reasons several health-promoting properties have been attributed to the microorganisms of this genus and increasing bifidobacterial levels in the GIT is often a treatment target of dietary intervention strategies. So far, there have been no reports of sepsis related to bifidobacteria use in otherwise healthy individuals. Therefore, the current evidence strongly suggests that bifidobacteria are extremely safe to use in the general population. In fact, the most commonly used Bifidobacterium species are considered to be generally safe (GRAS), and in Europe such microorganisms have been granted a Qualified Presumption of Safety (QPS) status.46 Despite the widespread use of the

genus Bifidobacterium, in particular the species B. animalis subsp. lactis, in commercial probiotic products, these probiotics have never been related to sepsis associated with probiotic use, demonstrating the low pathogenic potential of bifidobacteria. This extensive use of bifidobacteria as probiotic agents, together with the numerous clinical trials conducted with these microorganisms, constitutes good proof of safety. As of July 2009, 43 undergoing or completed human clinical trials in which bifidobacteria were administered can be found registered at ClinicalTrials.gov. During last year alone, more than 25 human intervention studies, in which around 2000 volunteers—including healthy adults and elderly, but also full-term and preterm infants, HIV infected children, allergic subjects, pregnant women or IBS patients among others— received a test product containing bifidobacteria, have been published without reporting any detrimental effects. In vitro assessments offer means to investigate the safety of bifidobacteria based on the intrinsic properties of the strains. Several different in vitro approaches have been used in the safety assessment of bifidobacteria. In vitro tests assessing the resistance to antibiotics and the presence of mobile antibiotic resistance genes are common and constitute perhaps the major safety concern regarding bifidobacteria. Several studies have attempted to identify the presence of relevant virulence determinants in bifidobacteria. However, to date such determinants have not been identified. It should be noted, however, that classical risk assessments, which are used for pathogens, might not always be directly applicable to microorganisms such as bifidobacteria, which become members of

50

the normal healthy intestinal microbiota soon after birth and are also components of normal human diet. In pathogens, pathogenicity is normally a consequence of several properties of the strain. The possible presence of such a property in a strain of low infective potential and low clinical significance does not imply that the strain is pathogenic or poses a risk to health.47 An example of this is the ability to adhere to human mucosa, which is a virulence factor in the case of true pathogens, but is also an essential feature of many commensal microbes with very low pathogenic potential, such as bifidobacteria. To date, no clear virulence factors similar to those associated with pathogenic microorganisms have been identified for bifidobacteria.48 This is likely to result from the minimal infectivity of the bifidobacteria. Animal models have been also used in the safety ­evaluation of Bifidobacterium.49,50 Acute oral toxicity tests have been carried out without observing toxicity.49,51 A number of ­studies have examined the effects of bifidobacteria in ­experimentally induced colitis in murine models. Despite disruption of the intestinal barrier in colitic animals, side effects due to increased bacterial translocation seem not to be apparent. In contrast, improvement of clinical condition by reducing the inflammatory colitis of these animals has been often reported. Immunocompromised animal models of both adult and young animals have also been tested to evaluate safety without observing detrimental effects. However, many of the animal models applied in the safety assessment of probiotics were originally developed to study pathogenic microorganisms in which virulence traits are present, and therefore they may not be optimal for studying translocation of non-virulent microorganisms such as bifidobacteria. With regard to antibiotic resistance, Bifidobacterium ­species are resistant to aminoglycosides, metronidazole, and gram-negative spectrum antibiotics. They are also intrinsically resistant to mupirocin. In contrast, bifidobacteria are very susceptible to macrolides/lincosamides, vancomycin, rifampicin, spectinomycin, chloramphenicol, and β-lactams. The susceptibility to tetracyclines and cephalosporins varies among strains.52 One of the current major concerns for the safe use of Bifidobacterium strains is the presence of tetracycline resistance genes, especially tet(W), although other genes, such as tet(M), tet(O), tet(L), tet(W/32/O), and tet(O/W), have been detected, albeit much less frequently.53 In bifidobacteria, tet genes seem to be integrated in the chromosome, and they have not been associated with transposons or plasmids, but they are very often flanked by putative transposase genes.53–55 Transposases are enzymes that catalyze the movement of DNA segments among different locations by recognizing insertion sequences in the DNA, and they are thought to be involved in the mobilization of tet(W) genes in bifidobacteria.54 The long history of safe consumption and the lack of toxicity and virulence determinants support the safety of Bifidobacterium. On the other hand, the low incidence rate of infections due to these microorganisms seems to support this hypothesis. However, it is important to underline that there is no “zero risk” in microbiology and safety assessment is always essential. In fact, although very rare,

Molecular Detection of Human Bacterial Pathogens

some bifidobacteria, probably of intestinal origin, have been ­associated with a few cases of bacteremia, usually in patients with underlying diseases.56 Reduced immune function is the main common denominator in these patients and such patients are prone to bacteremia in general and not ­bifidobacterial ­infection in particular. Of the about 2000 culture-positive cases of cervicofacial actinomycoses assessed by Pulverer et al., none was due to bifidobacteria.57 The few cases of infection or sepsis reported have been mainly related to Bifidobacterium eriksonii (also known as Actinomyces eriksonii) which has been considered a synonym of B. dentium.58 Often these cases are related with subjects with subjacent problems and correspond to ­polymicrobial infections.59 A fatal case of pulmonary infection due to B. eriksonii in a 52-year-old alcoholic patient with ­periodontal disease was reported.60 The presence of bifidobacteria was also observed in a case of infected cavitating lung tumor61 and a case of pericarditis.62 B. adolescentis was isolated in a case of urinary tract infection in a child.63 The presence of B. dentium (named as A. eriksonii), often together with other microorganisms, in some rare cases of abscesses has also been observed.64,65 B. breve was reported to be the causative agent of a neonatal meningitis case, being the second case of bifidobacterial meningitis reported to date.66 Ha et al.67 published a rare case of B. longum septicemia. The patient had undergone acupuncture for a lumbar hernia and small needles had been inserted in the lumbar area and left there. Other than the herniated intervertebral disk the subject was healthy, thus it seems that needle contamination or intestinal punctures by a needle were the likely causes of the infection. Despite these few and isolated cases, the pathogenicity of bifidobacteria appears to be very low. Bourne et al.68 carried out over 90,000 blood cultures from subjects with bacteremia symptoms, recovering over 9000 bacterial isolates. Only 10 out of those isolates were bifidobacteria, and in five cases it was possible to identify the strain as B. eriksonii. All these isolates were obtained from nine patients (in two of the cases, other microorganisms were also found) with underlying conditions (pregnancy complications, gastrointestinal disorders, systemic lupus). To date, the major concern regards caries and mainly B. dentium and Bifidobacterium-related genera such as Parascardovia and Scardovia, as these microorganisms may be found forming part of the complex microbial community present in caries.69,70 Clinical trials using bifidobacteria, along with the ­widespread and safe use of this microbial genus worldwide, constitute the most compelling evidence of the safety of probiotics. Although very often the main target outcome in these intervention trials focuses on the health benefits of the strain, the safety of the administration and the potential adverse events, if any, are often reported as a secondary outcome. Clinical safety trials enable the in vivo evaluation of the effects in humans in a controlled manner, with a special focus on the attributes relevant to the safety of the administered strain and the factors contributing to possible adverse

51

Bifidobacterium

events. However, these clinical investigations are most commonly carried out with healthy volunteers. Considering the infection cases observed, it is clear that safety of bifidobacteria may be of particular interest to specific groups, such as neonates, who have compromised immune systems, or diseased patients. In neonates and low-birth weight infants, successful clinical interventions have been carried out,71 and clinical trials suggest that these microorganisms are safe to use in follow-on formulas and growing-up milks.72 Clinical evaluation in elderly populations is also of special interest, since elderly subjects commonly have health-related problems including infections and gastrointestinal problems. The safety and the lack of adverse events following the consumption of Bifidobacterium strains by elderly subjects have also been demonstrated. However, the possible risks may be elevated in certain high-risk populations, and the demonstration of the safety of a certain strain in the general population does not necessarily imply that the administration of the same strain is equally safe in high-risk populations. In fact, despite the overwhelming evidence supporting bifidobacterial safety, there may be potentially adverse events, as demonstrated by a recent study carried out in severely ill patients with acute pancreatitis. Besselink et al.73 investigated the effects of a probiotic mix containing Lactobacillus, Lactococcus, and Bifidobacterium strains in a randomized placebo-controlled trial that included 296 adults with a first episode of high-risk acute pancreatitis. The probiotic mix was administered by nasojejunal feeding tube, and had been specifically designed, on the basis of previous studies, to inhibit the growth of pathogens important in pancreatic necrosis.74 The authors found a 2.53-fold increase in mortality risk in probiotic-treated participants, and nine cases of bowel ischemia (eight fatal) in the probiotic group, with no bowel ischemia in the placebo group. It is known that small bowel ischemia, increased intestinal permeability, and increased bacterial translocation are all associated with acute pancreatitis. In these cases, the direct application of probiotic bacteria to an already damaged small intestinal mucosa may have precipitated a local inflammatory response leading to increased risk of small bowel ischemia. However, it is not known why exactly mortality was higher in the treatment group, and although the mortality was associated with randomization, this does not necessarily imply that the probiotic itself was the causative factor.75 In these cases, the gastrointestinal system plays a central role in the multisystem organ failure, leading to a breakdown of the intestinal barrier that increases translocation of bacteria and their components into the systemic circulation. In spite of that, translocation is difficult to induce in healthy humans, and if this occurs, detrimental effects have not been demonstrated for bifidobacteria. Due to the general consideration as safe of bifidobacteria, most cases of the presence of these microorganisms in infection sites have been considered as contaminants and, much less probably, as pathogens.76 In any case, despite the excellent overall safety record of bifidobacteria, they should be used with caution in certain specific patient groups with severe underlying diseases and reduced immune function. It

is important to note that such patients in general have a high risk for bacteremia and, very often, intestinal bacteria are the origin of the infection.

5.1.4  Diagnosis Other than the infection cases reported, bifidobacteria may have a profound effect on human health. Low levels of intestinal bifidobacteria population and the misbalance of intestinal microbiota are thought to play an important role in the etiology and pathogenesis of different inflammatory, metabolic, and autoimmune diseases. Thus, alterations in intestinal microbiota composition are found in patients with allergies or those suffering from different health disorders.42,43,77,78 Whether these alterations on the microbiota composition and levels are the cause or consequence of illness is currently far from known. In any case, therapy with bifidobacteria and other probiotics have been successfully used for the alleviation of symptoms of these chronic disorders.79,80 The gut, particularly the colon, is largely the organ containing the highest population and diversity of Bifido­ bacterium species. Owing to this fact, methods for detection and enumeration of these microorganisms have mainly been developed focusing on intestinal bifidobacteria, taking into account the difficulty of isolation and characterization inherent in the high number and diversity of other microorganisms sharing the same environment. These methods have also been subsequently adapted and applied to the study of bifidobacteria in other locations. In this respect, it is worth ­mentioning the recent studies on Bifidobacterium species present in breast milk and the possible transmission of bifidobacterial populations, or their DNA, from mothers to children through ­placenta and breastfeeding.21,81,82 As deduced from that indicated above, although bifidobacteria have not been directly considered as the causative agent of any pathology, the development of quantitative and qualitative methods of detection and identification of these microorganisms has, and will, greatly contribute to gain insight into the relationship between levels and species composition of bifidobacteria populations and the health status of the host. The correct identification and typification of Bifido­ bacterium strains is the starting point to establish its microbiological properties and safety. Thus, it is necessary to use adequate tools to provide proper strain identification, to track Bifidobacterium strains during food production, during their GIT passage and establishment, or to identify potential clinical isolates. 5.1.4.1  Phenotypic Techniques The traditional identification of bifidobacteria has been based on phenotypic features. Cell morphology, production of metabolites, enzymatic activities and sugar fermentation profiles are the most commonly analyzed phenotypic characteristics. The association of a typical branched shape with the presence of fructose-6-phosphate phosphoketolase (F6PPK) activity is the first indication that a strain belongs to the genus Bifidobacterium.83 However, not all strains have

52

a V or clamped shape, and the levels of F6PPK activity vary enormously among species and strains.84 Furthermore, the identification at species/strain level have some additional problems, and the classical phenotyping, such as sugar fermentation profiles, transaldolase serotyping, cell wall composition, and the study of the F6PPK isoforms, is clearly not discriminative enough to reach species, subspecies, and biotype-level identification with accuracy. In fact, most cases of bifidobacteria misidentifications are due to inappropriate phenotypic methods.85 Several different quantitative and qualitative methodologies have been used for Bifidobacterium microbiota assessment.86,87 Traditional methods for the study of qualitative and quantitative composition of intestinal bifidobacteria have been carried out by the cultivation of feces. In some cases, it has also been considered that there are different mucosa-associated intestinal bacteria that can differ from feces in biopsies or samples taken from healthy individuals88 or patients.43,89 Due to the anaerobic nature of bifidobacteria, the samples must be handled in anaerobic conditions and processed immediately or shortly after collection. Bifidobacteria are nutritionally demanding microorganisms and therefore are cultured on suitable complex growth media (such as MRS, Rogosa, Beerens, and trypticase phytone yeast extract). Both nonselective and selective differential media have been used for growth and counting, with and without the addition of selective agents.90–92 The choice of one medium or another is dependent on the other bacteria present in the sample and on the method used for subsequent identification. Besides the characteristic “bifid” cell morphology and the F6PPK activity of the genus Bifidobacterium, another phenotypic method for identification at the species level is based on the determination of the cellular fatty acid composition93 with the help of the chromatographic MIDI system directly in feces42 or in previously isolated cultures.92 In spite of the accuracy of the method, it is important to mention that the composition of fatty acids changes notably in bifidobacteria stressed cells,94 which can lead to misidentifications if other alternative methods were not used in those cases. The difficulty of cultivation of anaerobic intestinal microorganisms meant that until a few years ago, their study was restricted to the cultivable species and among them, only to the viable population. This led to the overestimation of some species and the underestimation of others. In addition, not all Bifidobacterium species are recovered from selective media with the same efficiency,90,91 which could draw a distorted picture of the bifidobacteria population inhabiting complex human ecosystems. 5.1.4.2  Molecular Techniques In the last few years, new molecular tools have been developed and applied, as alternative or complementary methods to traditional cultures, for the identification and typing of bifidobacteria. Most of these techniques use 16S rRNA and its encoding genes as target molecules, on which specific PCR primers and hybridization probes are designed to detect particular species or groups of microorganisms, including

Molecular Detection of Human Bacterial Pathogens

the genus Bifidobacterium95–98 and some of its species.40,96,97 Moreover, the use of 16S rRNA instead of the DNA (by obtaining cDNA from mRNA through reverse-transcriptase PCR reactions) can provide data on the activity and viability of microorganisms in their ecosystem rather than on cellular levels as occurs when DNA is the target molecule. Among the most extensively used molecular tools for qualitative assessment of bifidobacteria is the sequence analysis of amplified 16S RNA genes, by PCR amplification from biological samples, followed by cloning and sequencing of the amplified DNA. Other widely used techniques are TGGE and DGGE, using primers for PCR amplification with one of these primers being tagged to a GC clamp to avoid complete dissociation of DNA strands. Amplified fragments are subsequently separated by denaturing gel electrophoresis through a ­gradient of ­temperature (TGGE) or denaturant agent (DGGE).99 Furthermore, PCR amplification products can be separated by high-performance liquid chromatography (HPLC).100 Analysis of terminal-restriction fragment length polymorphism (T-RFLP) by digestion of fluorescent labeled fragments with suitable restriction enzymes101 or the use of Oligonucleotide arrays102 may be also utilized. Sequencing of ribosomal rRNA genes (rDNA) is nowadays the standard methodology for bifidobacteria identification. The bacterial rRNA genes are organized in rrn operons containing the genes for the three ribonucleic acids (5S RNA, 16S RNA, and 23S RNA), with bifidobacteria harboring from two to six operons, depending on the species.10,30 The 16S and 23S rRNA genes, as well as the 16S23S spacer region (Intergenic Transcribed Sequence, ITS), showed nucleotide fingerprints with different discriminatory levels. Sequencing of the 16S rRNA can be used for identification purposes or to establish phylogenetic relationships among species, subspecies, and biotypes,5,97,103 whereas the 16S-23S rRNA ITS sequence is much more variable than the 16S rRNA structural gene, even within closely related taxa, which makes it a valuable target for species identification by using species-specific primers,104 but also its higher discriminatory capacity may allow the differentiation of strains.105 Recently, multilocus sequence typing (MLST) methods have been applied for the typification of Bifidobacterium species and strains. MLST methods made use of a DNA sequencing procedure to characterize different housekeeping  gene loci in the bacterial genome.106 Its application in bifidobacteria has been shown to be highly discriminatory, the target genes studied for detailed identification and classification purposes down to subspecies level being numerous.5,107–110 Application of a variety of PCR protocols has enabled the differentiation between strains of the same species. Random amplification of polymorphic DNA (RAPD) techniques, amplified ribosomal RNA restriction analysis (ARDRA), ERIC (Enterobacterial Repetitive Intergenic Consensus sequence)-PCR, and REP (Repetitive Extrogenic Palindromic)-PCR are the most common PCR-based methods for bifidobacterial typing.4,14,111–113 Other PCR-based

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Bifidobacterium

typing approaches include T/DGGE (temperature/denaturant gradient gel electrophoresis), amplified fragment length polymorphism (AFLP), PCR coupled to enzyme-linked immunosorbent assay (ELISA), triplicate arbitrarily primer (TAP)-PCR, restriction fragment length polymorphism (RFLP)-PCR, and multiplex PCR.108,111,114–116 Some other methods for bifidobacteria identification and typing used DNA profiling, such as plasmid analysis, RFLP, or PFGE (pulsed field gel electrophoresis) of total DNA.4,111,117 PFGE is often considered the best technique for strain-­specific typification and has shown a high discriminatory power. Among the nucleic acids hybridization methods, ribotyping, microplate blot, and microarray analysis have also been used for bifidobacterial typing.102,111,118 Finally, it is worthwhile to point out that other methods based on metabolite ­production, have also been applied to the identification and typing of Bifidobacterium such as chromatographic analysis of organic acids or the analysis of the intrinsic fluorescence of aromatic amino acids.119,120 Qualitative techniques provide a picture of the most abundant species and the diversity in microbial ecosystems but fail to provide data on the relative abundance of microbial groups as well as on the less abundant species. Among the quantitative techniques most extensively used for Bifidobacterium is fluorescent in situ hybridization (FISH), with specific probes coupled to flow cytometry or fluorescence microscopy detection. Although this technique has widely been used to study the evolution of intestinal populations,45,95,121 it is extremely laborious and time consuming, and is influenced by cell permeability or the ribosome content of cells. Quantitative real-time PCR is a faster tool to study complex microbial communities. Different PCR assays have been developed for fecal bifidobacteria and its species, or focused on the metabolically active microorganisms targeting 16S rDNA or rRNA, respectively, by using probes labeled with different fluorescent dyes. “Omics” (metagenomics, metaproteomics, and metabolomics) are very recent, high throughput, and promising techniques that can also be applied to the study of bifidobacteria in complex ecosystems. The sensitivity level of genetic methods for identification and typing of bacterial isolates is obviously limited by the previous sensitivity of the culture media and conditions used for isolating the microorganisms.

5.2  METHODS 5.2.1  Sample Preparation Most of the studies evaluating bifidobacterial population ­composition and levels have used fecal samples as starting material. However, the same methods with slight modifications can be used to determine the presence or levels of ­bifidobacteria in other matrixes such as human tissues or fluids. Both culture-dependent and culture-independent methods are available to assess Bifidobacterium populations. Depending on the method used for the detection of the microorganism, different sample preparation procedures are needed.

When a culture-dependent method is going to be used, isolation of the microorganisms is required prior to identification by phenotypic or genotypic techniques. Given the complex microbial community present in fecal samples, isolation of bifidobacteria is a challenging issue. Typically, the fecal sample is diluted in an isotonic buffer (e.g., PBS) and homogenized using a blender. Decimal dilutions are then prepared and 0.1 mL of the appropriate dilution is plated in an adequate selective or differential culture media.91,92 After the incubation of the plates at 37°C for 48 h under anaerobic conditions in an anaerobic chamber or a jar with an anaerobiosis system (e.g., Anaerocult A, Merck), putative bifidobacterial colonies may be picked up and their identity further confirmed by phenotypic or molecular techniques as stated in Section 5.1.4. Very often, molecular methods of identification focus on DNA analyses that require a step of extraction of DNA from the studied strain. Bifidobacteria are not easily lysed, and for this reason lysis buffers are often supplemented with lysozyme and mutanolysin to improve the lysis efficiency, thereby increasing the yield of the DNA extraction. Nowadays, in addition to the traditional methods of DNA extraction using phenol/chloroform, a number of DNA extraction kits are commercially available from the main molecular biology suppliers and are very often used for bacterial DNA extraction. Culture-independent methods do not require a previous bacterial isolation step. In this case, the starting material may be a complex sample (i.e., fecal sample). The total DNA in the sample is extracted previously to the specific detection of bifidobacterial DNA in the complex DNA pool extract. Different DNA extraction methods for such complex samples have been reported in the literature19,99 and commercial kits are also available (e.g., QIAamp DNA Stool Kit, Qiagen) to this end.

5.2.2  Detection Procedures Among the easiest and most currently used molecular tools to detect the presence of bifidobacteria in complex samples is the use of PCR with specific primers. Several specific primers have been designed for the genus Bifidobacterium97,98 and for different bifidobacterial species.40,96,97,122 These primers have been extensively used for the culture-independent detection of bifidobacteria, mainly in fecal samples but also in other matrixes, including food products. Procedure 1. Prepare the PCR mix (most commonly 25 or 50 µL) containing PCR buffer, DNA polymerase, dNTPs, and the appropriate primers (genus or species-specific depending on the aim of the study). Add DNA extract (most often 2–5 µL) or an equal amount of water (negative control). 2. Run the PCR program using the optimal conditions for the DNA polymerase used and the adequate annealing temperature (depending in the primers used).

54



Molecular Detection of Human Bacterial Pathogens

3. After completion of PCR, add DNA loading buffer and visualize amplification products by standard agarose gel electrophoresis and ethidium bromide staining.

Note: Some of these primers have also been used for ­determination of bifidobacterial levels by means of quantitative real-time PCR. Two approaches may be used to this end; a combination of the primers with a DNA dying agent such as SYBR Green (SYBR Green assay),15,21,44 or the combined use of the primers with a labeled internal oligonucleotide probe (5´nuclease assay).40,98,122

5.3  CONCLUSION AND FUTURE PERSPECTIVES Bifidobacteria are normal inhabitants of the human ­intestinal tract, constituting an important part of the microbiota of the colon, where they reach very high numbers. These ­microorganisms have shown a high adaptation to their ­environment and are fastidious and difficult to cultivate in the laboratory. However, the development of techniques to grow anaerobic microorganisms, together with the recent ­development of culture-independent methods, has allowed an improvement of our knowledge on Bifidobacterium ­physiology and ecology. Although some rare cases of infection have been reported, usually in subjects with underlying diseases, the major ­concern about Bifidobacterium safety regards the potential role of B.  dentium in dental caries. In fact, these ­microorganisms have shown a very low pathogenic potential, as has been demonstrated by in vitro animal and human studies. Moreover, several studies have indicated that reduced intestinal bifidobacterial levels may be related to different diseases and ­several ­health-promoting properties have been attributed to these microorganisms. For these reasons, increasing bifidobacterial levels has often been taken as a treatment target. Different nutritional intervention strategies have been developed to this end, mainly using probiotic bacteria, including bifidobacteria, and prebiotic compounds. In this regard, it is important to underline that there is no “zero risk” and that the addition of any microorganisms to products for human or ­animal ­consumption requires a rigorous assessment of its safety. In the near future, the recent developments on ­molecular methods and the availability of bifidobacterial genomes will improve our understanding of the biology of the genus Bifidobacterium, the interaction with the host, and the role in health and disease.





REFERENCES



1. Scardovi, V., Genus Bifidobacterium. Orla-Jensen 1924, 472AL, In Sneath, P.H.A., Mair, N.S., Sharpe, M.E., and Holt, J.G. (Editors), Bergey’s Manual of Systematic Bacteriology, Vol. 2, pp. 1418–1434, Williams and Wilkins, Baltimore, USA, 1986. 2. Simpson, P.J. et al., Intrinsic tolerance of Bifidobacterium species to heat and oxygen and survival following spray drying and storage, J. Appl. Microbiol., 99, 493, 2005.



3. Tissier, M.H., Réchérches sur la flore intestinale normale et pathologique du nourisson (thesis), University of Paris, France, 1900. 4. Satokari, R.M. et al., Molecular approaches for the detection and identification of bifidobacteria and lactobacilli in the human gastrointestinal tract, Syst. Appl. Microbiol., 26, 572, 2003. 5. Ventura, M. et al., Analysis of bifidobacterial evolution using a multilocus approach, Int. J. Syst. Evol. Microbiol., 56, 2783, 2006. 6. Ventura, M. et al., Genomics of Actinobacteria: tracing the evolutionary history of an ancient phylum, Microbiol. Mol. Biol. Rev., 71, 495, 2007. 7. Okamoto, M. et al., Bifidobacterium tsurumiense sp. nov., from hamster dental plaque, Int. J. Syst. Evol. Microbiol., 58, 144, 2008. 8. Killer, J. et al., Bifidobacterium bombi sp. nov., from the ­bumblebee digestive tract, Int. J. Syst. Evol. Microbiol., 59, 2020, 2009. 9. Watanabe, K. et al., Bifidobacterium mongoliense sp. nov., from airag, a traditional fermented mare’s milk product from Mongolia, Int. J. Syst. Evol. Microbiol., 59, 1535, 2009. 10. Ventura, M. et al., Insights into the taxonomy, genetics and physiology of bifidobacteria, Antonie Van Leeuwenhoek, 86, 205, 2004. 11. Felis, G.E., and Dellaglio, F., Taxonomy of lactobacilli and bifidobacteria, Curr. Issues Intest. Microbiol., 8, 44, 2007. 12. Masco, L. et al., Culture-dependent and culture-independent qualitative analysis of probiotic products claimed to contain bifidobacteria, Int. J. Food Microbiol., 102, 221, 2005. 13. Reuter, G., The Lactobacillus and Bifidobacterium microflora of the human intestine: composition and succession, Curr. Issues Intest. Microbiol., 2, 43, 2001. 14. Turroni, F. et al., Exploring the diversity of the bifidobacterial population in the human intestinal tract, Appl. Environ. Microbiol., 75, 1534, 2009. 15. Turroni, F. et al., Microbiomic analysis of the bifidobacterial population in the human distal gut, ISME J., 3, 745, 2009. 16. Guarner, F., and Malagelada, J-R., Gut flora in health and disease, Lancet, 381, 512, 2003. 17. Bäckhed, F. et al., Host-bacterial mutualism in the human intestine, Science, 307, 1915, 2005. 18. Isolauri, E., Salminen, S., and Ouwehand, A.C., Probiotics, Best Pract. Res. Clin. Gastroenterol., 18, 299, 2004. 19. Eckburg, P.B. et al., Diversity of the human intestinal microbial flora, Science, 308, 1635, 2005. 20. Beighton, D. et al., Isolation and identification of Bifido­ bacteriaceae from human saliva, Appl. Environ. Microbiol., 74, 6457, 2008. 21. Martin, R. et al., Isolation of bifidobacteria from breast milk and assessment of the bifidobacterial population by PCRdenaturing gradient gel electrophoresis and quantitative realtime PCR, Appl. Environ. Microbiol., 75, 965, 2009. 22. Lamendella, R. et al., Bifidobacteria in feces and environmental waters, Appl. Environ. Microbiol., 74, 575, 2008. 23. Masco, L. et al., In vitro assessment of the gastrointestinal transit tolerance of taxonomic reference strains from human origin and probiotic product isolates of Bifidobacterium, J. Dairy Sci., 90, 3572, 2007. 24. Cotter, P.D., and Hill, C., Surviving the acid test: Responses of gram-positive bacteria to low pH, Microbiol. Mol. Biol. Rev., 67, 429, 2003. 25. Matsumoto, M., Ohishi, H., and Benno, Y., H+-ATPase activity in Bifidobacterium with special reference to acid tolerance, Int. J. Food Microbiol., 93, 109, 2004.

Bifidobacterium 26. Begley, M., Gahan, C.G., and Hill, C., The interaction between bacteria and bile, FEMS Microbiol. Rev., 29, 625, 2005. 27. Scardovi, V., and Trovatelli, L.D., The fructose-6-phosphate shunt as peculiar pattern of hexose degradation in the genus Bifidobacterium, Ann. Microbiol. Enzimol., 15, 19, 1965. 28. Sánchez, B. et al., Bacterial and eukaryotic phosphoketolases: Phylogeny, distribution and evolution, J. Mol. Microbiol. Biotechnol., 18, 37, 2010. 29. Meile, L. et al., Characterization of the D-xylulose 5-phosphate/ D-fructose 6-phosphate phosphoketolase gene (xfp) from Bifidobacterium lactis, J. Bacteriol., 183, 2929, 2001. 30. Schell, M.A. et al., The genome sequence of Bifidobacterium longum reflects its adaptation to the human gastrointestinal tract, Proc. Natl. Acad. Sci. USA, 99, 14422, 2002. 31. Sela, D.A. et al., The genome sequence of Bifidobacterium longum subsp. infantis reveals adaptations for milk utilization within the infant microbiome, Proc. Natl. Acad. Sci. USA, 105, 18964, 2008. 32. Kim, J.F. et al., Genome sequence of the probiotic bacterium Bifidobacterium animalis subsp. lactis AD011, J. Bacteriol., 191, 678, 2009. 33. Ryan, S.M., Fitzgerald, G.F., and van Sinderen, D., Transcriptional regulation and characterization of a novel beta-fructofuranosidase-encoding gene from Bifidobacterium breve UCC2003, Appl. Environ. Microbiol., 71, 3475, 2005. 34. McKellar, R.C., and Modler, H.W., Metabolism of fructooligosaccharides by Bifidobacterium spp., Appl. Microbiol. Biotechnol., 31, 537, 1989. 35. Gibson, G.R., and Roberfroid, M.B., Dietary modulation of the human colonic microbiota: Introducing the concept of prebiotics, J. Nutr., 125, 1401, 1995. 36. Kurokawa, K. et al., Comparative metagenomics revealed commonly enriched gene sets in human gut microbiome, DNA Res., 14, 169, 2007. 37. Klijn, A., Mercenier, A., and Arigoni, F., Lessons from the genomes of bifidobacteria, FEMS Microbiol. Rev., 29, 491, 2005. 38. Barrangou, R. et al., Comparison of the complete genome sequences of Bifidobacterium animalis subsp. lactis DSM10140 and B1–04, J. Bacteriol., 191, 4144, 2009. 39. Ventura, M. et al., Genome-scale analices of health promoting bacteria: Probiogenomics, Nat. Rev. Microbiol., 7, 61, 2009. 40. Gueimonde, M. et al., Quantitative assessment of faecal ­bifidobacterial populations by using novel real-time PCR assays, J. Appl. Microbiol., 102, 1116, 2007. 41. Favier, C. et al., Fecal beta-D-galactosidase production and Bifidobacteria are decreased in Crohn’s disease, Dig. Dis. Sci., 42, 817, 1997. 42. Kalliomäki, M. et al., Distinct patterns of neonatal gut ­microflora in infants in whom atopy was and was not developing, J. Allergy Clin. Immunol., 107, 129, 2001. 43. Gueimonde, M. et al., Qualitative and quantitative analyses of the bifidobacterial microbiota in the colonic mucosa of patients with colorectal cancer, diverticulitis and inflammatory bowel disease, World. J. Gastroenterol., 13, 3985, 2007. 44. Collado, M.C. et al., Imbalances in faecal and duodenal Bifidobacterium species composition in active and non-active coeliac disease, BMC Microbiol., 8, 232, 2008. 45. Kerckhoffs, A.P. et al., Lower Bifidobacteria counts in both duodenal mucosa-associated and fecal microbiota in irritable bowel syndrome patients, World J. Gastroenterol., 15, 2887, 2009.

55 46. EFSA, European Food Safety Authority Scientific Committee (EFSA) public consultation on the Qualified Presumption of Safety (QPS) approach for the safety assessment of microorganisms deliberately added to food and feed. Annex 3: Assessment of gram positive non-sporulating bacteria with respect to a qualified presumption of safety. http://www.efsa. europa.eu/en/science/sc_commitee/sc_consultations/sc_consultation_qps.html. 2007. 47. Borriello, S.P. et al., Safety of probiotics that contain lactobacilli or bifidobacteria, Clin. Infect. Dis., 36, 775, 2003. 48. Ouwehand, A.C., Saxelin, M., and Salminen, S., Assessment of potential risk factors and related properties of clinical, faecal and dairy Bifidobacterium isolates, Biosci. Microflora, 23, 37, 2004. 49. Zhou, J.S. et al., Acute oral toxicity and bacterial translocation studies on potentially probiotic strains of lactic acid bacteria, Food Chem. Toxicol., 38, 153, 2000. 50. Zhou, J.S. et al., Safety assessment of potential probiotic lactic acid bacterial strains Lactobacillus rhamnosus HN001, Lb. acidophilus HN017, and Bifidobacterium lactis HN019 in BALB/c mice, Int. J. Food Microbiol., 56, 87, 2000. 51. Kabeir, B.M. et al., Safety evaluation of Bifidobacterium pseudocatenulatum G4 as assessed in BALB/c mice, Lett. Appl. Microbiol., 46, 32, 2008. 52. Zhou, J.S. et al., Antibiotic susceptibility profiles of new ­probiotic Lactobacillus and Bifidobacterium strains, Int. J. Food Microbiol., 98, 211, 2005. 53. Florez, A.B. et al., Molecular analysis of tet(W) gene-mediated tetracycline resistance in dominant intestinal Bifidobacterium species from healthy humans, Appl. Environ. Microbiol., 72, 7377, 2006. 54. Kazimierczak, K.A., Flint, H.J., and Scott, K.P., Comparative analysis of sequences flanking tet(W) resistance genes in multiple species of gut bacteria, Antimicrob. Agents Chemother., 50, 2632, 2006. 55. van Hoek, A.H. et al., Resistance determinant erm(X) is borne by transposon Tn5432 in Bifidobacterium thermophilum and Bifidobacterium animalis subsp. lactis, Int. J. Antimicrob. Agents, 31, 544, 2008. 56. Gasser, F., Safety of lactic acid bacteria and their occurence in human clinical infection, Bull. Ins. Pasteur, 92, 45, 1994. 57. Pulverer, G., Schütt-Gerowitt, H., and Schall, K.P., Human cervicofacial actinomycoses: Microbiological data from 1997 cases, Clin. Infect. Dis., 37, 490, 2003. 58. Biavati, B., Scardovi, V., and Moore, W.E.C., Electrophoretic patterms of proteins in the genus Bifidobacterium and proposal of four new species, Int. J. Syst. Bacteriol., 32, 358, 1982. 59. Brook, I., Isolation of non-sporing anaerobic rods from infections in children, J. Med. Microbiol., 45, 21, 1996. 60. Green, S.L., Case-report: fatal anaerobic pulmonary infection due to Bifidobacterium eriksonii, Postgrad. Med., 63, 187, 1978. 61. Liao, W-Y. et al., Bacteriology of infected cavitating lung tumor, Am. J. Respir. Crit. Care Med., 161, 1750, 2000. 62. Brook, I., Pericarditis caused by anaerobic bacteria, Int. J. Antimicrob. Agents, 33, 297, 2009. 63. Brook, I., Urinary tract infection caused by anaerobic bacteria in children, Urology, 16, 596, 1980. 64. Cryan, B. et al., Epidural abscess due to streptococcus milleri and Bifidobacterium species, J. Infect., 23, 214, 1991.

56 65. Civen, R., Vaisanen, M.L., and Finegold, S.M., Peritonsilar abscess, retropharyngeal adscess, mediastinitis, and nonclostridial anaerobic myonecrosis: A case report, Clin. Infect. Dis., 16, S299, 1993. 66. Nakazawa, T. et al., Neonatal meningitis caused by Bifidobacterium breve, Brain Dev., 18, 160, 1996. 67. Ha, G.Y. et al., Case of sepsis caused by Bifidobacterium longum, J. Clin. Microbiol., 37, 1227, 1999. 68. Bourne, K.A. et al., Bacteremia due to Bifidobacterium, Eubacterium or Lactobacillus; twenty-one cases and review of the literature, Yale J. Biol. Med., 51, 505, 1978. 69. Chhour, K-L. et al., Molecular analysis of microbial diversity in advanced caries, J. Clin. Microbiol., 43, 843, 2005. 70. Mantzourani, M., Fenlon, M., and Beighton, D., Association between bifidobacteriaceae and the clinical severity of root caries lesions, Oral Microbiol. Immunol., 24, 32, 2009. 71. Hoyos, A.B., Reduced incidence of necrotizing ­enterocolitis associated with enteral administration of Lactobacillus acidophilus and Bifidobacterium infantis to neonates in an intensive care unit, Int. J. Infect. Dis., 3, 197, 1999. 72. Haschke, F. et al., Clinical trials prove the safety and efficacy of the probiotic strain Bifidobacterium Bb12 in follow-up ­formula and growing-up milks, Monatsschr. Kinderheilkd., 1146, S26, 1998. 73. Besselink, M.G.H. et al., Probiotic prophylaxis in predicted severe acute pancreatitis: A randomized, double-blind, placebocontrolled trial, Lancet, 371, 651, 2008. 74. van Minnen, L.P. et al., Modification of intestinal flora with multispecies probiotics reduces bacterial translocation and improves clinical course in a rat model of acute pancreatitis, Surgery, 141, 470, 2007. 75. Reid, G. et al., Probiotic prophylaxis in predicted severe acute pancreatitis, Lancet, 372, 112, 2008. 76. Liong, M.T., Safety of probiotics: translocation and infection, Nutr. Rev., 66, 192, 2008. 77. Sanz, Y. et al., Differences in faecal bacterial communities in coeliac and healthy children as detected by PCR and denaturing gradient gel electrophoresis, FEMS Immunol. Med. Microbiol., 51, 562, 2007. 78. Swidsinski, A. et al., Active Crohn’s disease and ulcerative colitis can be specifically diagnosed and monitored based on the biostructure of the fecal flora. Inflamm. Bowel Dis., 14, 147, 2008. 79. Picard, C. et al., Review article: Bifidobacteria ans probiotic agents-physiological effects and clinical benefits, Aliment. Pharmacol. Ther., 22, 495, 2005. 80. Zuccotti, G.V. et al., Probiotics in clinical practice: An ­overview, J. Int. Med. Res., 36, 1A, 2008. 81. Gueimonde, M. et al., Breast milk: A source of bifidobacteria for infant gut development and maturation? Neonatology, 92, 64, 2007. 82. Satokari, R. et al., Bifidobacterium and Lactobacillus DNA in the human placenta, Lett. Appl. Microbiol., 48, 8, 2009. 83. Ballongue, J., Bifidobacteria and probiotic action, In Salminen,  S., von Wright, A., and Ouwehand, A. (Editors), Lactic Acid Bacteria: Microbiological and Functional Aspects, 3rd ed., pp. 67–124S, Marcel Dekker, New York, 2004. 84. Orban, J.I., and Patterson, J.A., Modification of the phosphoketolase assay for rapid identification of bifidobacteria, J. Microbiol. Methods, 40, 221, 2000. 85. Huys, G. et al., Accuracy of species identity of commercial bacterial cultures intended for probiotic or nutritional use, Res. Microbiol., 157, 803, 2006.

Molecular Detection of Human Bacterial Pathogens 86. Gueimonde, M., and de los Reyes-Gavilán, C.G., Detection and enumeration of gastrointestinal microorganisms, In Lee, Y.K., and Salminen, S. (Editors), Handbook of Probiotics and Prebiotics, 2nd ed., pp. 25–43, John Wiley and Sons, Inc., New Jersey, USA, 2009. 87. Margolles, A., Mayo, B., and Ruas-Madiedo, P., Screening, identification, and characterization of Lactobacillus and Bifidobacterium strain. In Lee, Y.K., and Salminen, S. (Editors), Handbook of Probiotics and Prebiotics, 2nd ed., pp.  4–24, John Wiley and Sons, Inc., New Jersey, USA, 2009. 88. Delgado, S., Suárez, A., and Mayo, B., Bifidobacterial diversity determined by culturing and by 16S rDNA sequence analysis in feces and mucosa from ten healthy Spanish adults, Dig. Dis. Sci., 51, 1878, 2006. 89. Conte, M.P. et al., Gut-associated bacterial microbiota in paediatric patients with inflammatory bowel disease, Gut, 55, 1760, 2006. 90. Silvi, S., Rumney, C.J., and Rowland, I.R., An assessment of three selective media for bifidobacteria in faeces, J. Appl. Bacteriol., 81, 561, 1996. 91. Apajalahti, J.H.A. et al., Selective plating underestimates abundance and shows differential recovery of bifidobacterial species from human feces, Appl. Environ. Microbiol., 69, 5731, 2003. 92. Woodmansey, E. et al., Comparison of compositions and ­metabolic activities of fecal microbiotas in young adults and in antibiotic-treated and non-antibiotic treated elderly subjects, Appl. Environ. Microbiol., 70, 6113, 2004. 93. Eerola, E., and Lehtonen, O.P., Optimal data processing procedure for automatic bacterial identification by gas-liquid chromatography of cellular fatty acids. J. Clin. Microbiol., 26, 1745, 1998. 94. Ruiz, L. et al., Cell envelope changes in Bifidobacterium animalis ssp. lactis as a response to bile, FEMS Microbiol. Lett., 274, 316, 2007. 95. Langendijk, P.S. et al., Quantitative fluorescence in-situ hybridization of Bifidobacterium spp. with genus-specific 16S ribosomal-RNA-targeted probes and its application in fecal samples, Appl. Environ. Microbiol., 61, 3069, 1995. 96. Kaufmann, P. et al., Identification and quantification of Bifidobacterium species isolated from food with genus-specific 16S rRNA-targeted probes by colony hybridization and PCR, Appl. Environ. Microbiol., 63, 1268, 1997. 97. Matsuki, T., Watanabe, K., and Tanaka, R., Genus- and ­species-specific PCR primers for the detection and identification of bifidobacteria, Curr. Issues Intest. Microbiol., 4, 61, 2003. 98. Gueimonde, M. et al., New real-time quantitative PCR procedure for quantification of bifidobacteria in human fecal samples, Appl. Environ. Microbiol., 70, 4165, 2004. 99. Favier, C.F., de Vos, W.M., and Akkermans, A.D.L., Development of bacterial and bifidobacterial communities in feces of newborn babies, Anaerobe, 9, 219, 2003. 100. Goldenberg, O. et al., Molecular monitoring of the intestinal flora by denaturing high performance liquid chromatography, J. Microbiol. Meth., 68, 94, 2007. 101. Sakata, S. et al., Culture-independent analysis of fecal microbiota in infants, with special reference to Bifidobacterium species, FEMS Microbiol. Lett., 243, 417, 2005. 102. Boesten, R.J., Schuren, F.H., and de Vos, W.M., A Bifido­ bacterium mixed-species microarray for high resolution discrimination between intestinal bifidobacteria, J. Microbiol. Methods, 76, 269, 2009.

Bifidobacterium 103. Germond, J.E., Mamin, O., and Mollet, B., Species specific identification of nine human Bifidobacterium spp. in feces, Syst. Appl. Microbiol., 25, 536, 2002. 104. Gürtler, V., and Stanisish, V.A., New approaches to typing and identification of bacteria using the 16S-23S rDNA spacer region, Microbiol., 142, 3, 1996. 105. Leblond-Bourget, N. et al., 16S rRNA and 16S to 23S internal transcribed spacer sequence analyses reveal inter- and intraspecific Bifidobacterium phylogeny, Int. J. Syst. Bacteriol., 46, 102, 1996. 106. Maiden, M.C. et al., Multilocus sequence typing: A portable approach to the identification of clones within populations of pathogenic microorganisms, Proc. Natl. Acad. Sci. USA, 95, 3140, 1998. 107. Requena, T. et al., Identification, detection, and enumeration of human Bifidobacterium species by PCR targeting the transaldolase gene, Appl. Environ. Microbiol., 68, 2420, 2002. 108. Masco, L. et al., Polyphasic taxonomic analysis of Bifi­ dobacterium animalis and Bifidobacterium lactis reveals relatedness at the subspecies level: Reclassification of Bifido­ bacterium animalis as Bifidobacterium animalis subsp. animalis subsp. nov. and Bifidobacterium lactis as Bifidobacterium animalis subsp. lactis subsp. nov., Int. J. Syst. Evol. Microbiol., 54, 1137, 2004. 109. Yin, X. et al., The gene encoding xylulose-5-phosphate/­ fructose-6-phosphate phosphoketolase (xfp) is conserved among Bifidobacterium species within a more variable region of the genome and both are useful for strain identification, FEMS Microbiol. Lett., 246, 251, 2005. 110. Ventura, M. et al., Molecular characterization of hsp20, encoding a small heat shock protein of bifidobacterium breve UCC2003, Appl. Environ. Microbiol., 73, 4695, 2007. 111. Matto, J. et al., Genetic heterogeneity and functional properties of intestinal bifidobacteria, J. Appl. Microbiol., 97, 459, 2004. 112. Ventura, M. et al., Molecular microbial analysis of Bifido­ bacterium isolates from different environments by the

57 species-specific amplified ribosomal DNA restriction analysis (ARDRA), FEMS Microbiol. Ecol., 36, 113, 2001. 113. Krizova, J., Spanova, A., and Rittich, B., Evaluation of amplified ribosomal DNA restriction analysis (ARDRA) and species-specific PCR for identification of Bifidobacterium species, Syst. Appl. Microbiol., 29, 36, 2006. 114. Cusick, S.M., and O’Sullivan, D.J., Use of a single, triplicate arbitrarily primed-PCR procedure for molecular fingerprinting of lactic acid bacteria, Appl. Environ. Microbiol., 66, 2227, 2000. 115. Favier, C.F. et al., Molecular monitoring of succession of bacterial communities in human neonates, Appl. Environ. Microbiol., 68, 219, 2002. 116. Delcenserie, V. et al., Discrimination between Bifidobacterium species from human and animal origin by PCR-restriction fragment length polymorphism, J. Food Prot., 67, 1284, 2004. 117. Alvarez-Martín, P., Flórez, A.B., and Mayo, B., Screening for plasmids among human bifidobacteria species: Sequencing and analysis of pBC1 from Bifidobacterium catenulatum L48, Plasmid, 57, 165, 2007. 118. Wang, R.F. et al., DNA microarray analysis of predominant human intestinal bacteria in fecal samples, Mol. Cell. Probes, 18, 223, 2004. 119. Lee, K.Y., So, J.S., and Heo, T.R., Thin layer chromatographic determination of organic acids for rapid identification of ­bifidobacteria at genus level, J. Microbiol. Meth., 45, 1, 2001. 120. Ammor, M.S. et al., Reagentless identification of human ­bifidobacteria by intrinsic fluorescence, J. Microbiol. Meth., 69, 100, 2007. 121. Saulnier, D.M.A., Gibson, G.R., and Kolida, S., In vitro effects of selected synbiotics on the human faecal microbiota composition, FEMS Microbiol. Ecol., 66, 516, 2008. 122. Haarman, M., and Knol, J., Quantitative real-time PCR assays to identify and quantify fecal Bifidobacterium species in infants receiving a prebiotic infant formula, Appl. Environ. Microbiol., 71, 2318, 2005.

6 Corynebacterium Luis M. Mateos, Michal Letek, Almudena F. Villadangos, María Fiuza, Efrén Ordoñez, and José A. Gil

CONTENTS 6.1 Introduction........................................................................................................................................................................ 59 6.1.1 Classification, Morphology, and Biology............................................................................................................... 59 6.1.2 Phenotypic and Phylogenetic Characterization...................................................................................................... 60 6.1.2.1 Phenotypic Analysis................................................................................................................................ 60 6.1.2.2 Protein Analysis....................................................................................................................................... 60 6.1.2.3 Phylogenetic Analysis.............................................................................................................................. 64 6.1.3 Pathogenesis and Diagnosis.................................................................................................................................... 65 6.1.3.1 Corynebacterium Diphtheriae................................................................................................................ 65 6.1.3.2 Corynebacterium Species of the Lipophilic Group................................................................................. 66 6.1.3.3 Corynebacterium Species of the Nonlipophilic Group........................................................................... 67 6.2 Methods.............................................................................................................................................................................. 68 6.2.1 Sample Preparation................................................................................................................................................. 68 6.2.1.1 Culture Media for Corynebacterium....................................................................................................... 68 6.2.1.2 DNA Isolation.......................................................................................................................................... 68 6.2.1.3 RNA Isolation.......................................................................................................................................... 69 6.2.2 Detection Procedures.............................................................................................................................................. 69 6.2.2.1 Biolog (GP2) System................................................................................................................................ 69 6.2.2.2 API Coryne System................................................................................................................................. 69 6.2.2.3 Identifications Based on 16S rRNAs Genes............................................................................................ 70 6.3 Future Perspectives and Conclusions................................................................................................................................. 70 Acknowledgments........................................................................................................................................................................ 71 References.................................................................................................................................................................................... 71

6.1  INTRODUCTION 6.1.1  Classification, Morphology, and Biology The coryneform bacteria are a broad group of gram-positive, irregularly shaped rods (sometime oval- or club-shaped), aerobic or facultatively anaerobic, asporogenous, and nonpartially-acid-fast microorganisms classically used in the production of primary metabolites (amino acids, nucleotides, vitamins, etc.).1 The genus Corynebacterium was created in 1896 in order to assign a classification to Corynebacterium diphtheriae (the diphtheroid bacilli), as well as several animal pathogens and parasitic microorganisms.2 During most of the twentieth century, many microorganisms were included in this group based on their staining, morphological, and metabolic properties; in fact, in Bergey’s Manual of Determinative Bacteriology, the coryneform group comprised a broad and diverse number of bacteria of medical

(saprophytic animal and human pathogens), agricultural (plant pathogens), and industrial (saprophytic nonpathogenic amino-acid-producing strains) interest. In the 1990s, phylogenetic (based on 16S and 23S rDNA sequence analyses) and chemotaxonomic (cell wall composition and lipid profiles) analyses resulted in the separation of some members of the coryneform group from those corresponding to plant pathogens.3 Nonetheless, the number of species described for the genus Corynebacterium expanded, from 17 in the first edition of Bergey’s Manual of Systematic Bacteriology4 to more than 70 in the second edition (http://www.bergeys.org/). This large expansion of a bacterial genus is a rare occurrence in microbiology and reflects the enormous interest in these bacteria, mainly in medicine but also by industry. While very few data are available concerning the numbers of corynebacterial species present in nature, given what is known thus far about the genus Corynebacterium, it can be reasonably assumed 59

60

that corynebacteria are abundantly disseminated within a broad range of different natural habitats. In turn, much species diversity remains to be identified and characterized. The most prevalent human pathogen among the corynebacteria is C. diphtheriae, which continues to be a major cause of morbidity and mortality in undeveloped countries and increasingly in developing countries, where nontoxigenic members cause additional pathologies.5 Corynebacterium species other than those belonging to the C. diphtheriae group can be opportunistic or nosocomial pathogens of humans.6 Importantly, infections by these species are becoming increasingly common, mainly due to the remarkable increase in the number of susceptible individuals, especially patients with autoimmune disorders or undergoing immunosuppressive therapy as well as organ-transplanted patients. Species belonging to the genus Corynebacterium constitute a reasonably monophyletic group from the suborder Corynebacterineae, in the order Actinomycetales. They have a GC content of 51%–65% and an optimal temperature for growth of 28°C–37°C. Two broad groups of species from the genus can be distinguished based on their habitats and the consequences of the host-bacteria interaction: (i) Saprophytic members (nonpathogenic) are widely disseminated in different environments, such as soil, sea, dairy products, and animals. Of these, the main species used in industry is C. glutamicum. Additional species include C. ammoniagenes, C.  casei, C. callunae, C. efficiens, C. flavescens, and C. thermoaminogenes.7 (ii) Among the Corynebacterium species of medical importance, the best described is C. diphtheriae, but during the last few decades dozens of others have been recognized as emergent human and animal pathogens. Their optimal temperature for growth is 35°C–37°C and most are auxotrophic, with requirements for vitamins, amino acids, and/or nitrogen bases. Frequently, two broad subgroups of Corynebacterium members are distinguished based on an essential requirement for a particular nutrient, better growth in the presence of lipids (lipophilic group), or equivalent growth in the presence or absence of lipids (nonlipophilic group). Among the lipophilic group, species such as C. accolens, C. afermentans (lipophilum), C. appendicis, C. genitalium, C. hansenii, C. jeikeium, C. kroppenstedtii, C. lipophiloflavum, C. macginleyi, C. pseudogenitalium, C. resistens, C. sputi, C. tuberculostearicum, C. urealyticum, and C. ureicelerivorans are the most representative. Within the nonlipophilic group, a broader number of human pathogens (ascertained or potential) have been described, including C. diphtheriae, C. amycolatum, C. argentoratense, C. atypicum, C. auris, C. aurimucosum, C. confusum, C. coyleae, C. durum, C. freneyi, C. genitalum, C. glucuronolyticum, C. imitans, C. minutissimum, C. mucifaciens, C.  pilbarense, C.  pseudodiphtheriticum,

Molecular Detection of Human Bacterial Pathogens

C.  pseudotuberculosis, C.  resistens, C. riegelii, C.  simulans, C. singulare, C. striatum, C. sundvallense, C. thomssenii, C. timonense, C. tuscanie, C. ulcerans, and C. xerosis. In addition, the nonlipophilic group can be subdivided in fermentative and nonfermentative groups depending, respectively, on the ability or inability (to metabolize sugars under anoxic conditions). Table 6.1 summarizes most of the human-pathogenic Corynebacterium species isolated from clinical samples and characterized thus far, together with the most relevant information concerning these species.

6.1.2  Phenotypic and Phylogenetic Characterization 6.1.2.1  Phenotypic Analysis In-depth characterization of new isolates as members of Corynebacterium and their description as species of this genus rely on phenotypic analyses and thus on a combination of (i) morphological, (ii) physiological, and (iii) chemotaxonomic criteria. Preliminary identification has been traditionally achieved by Gram staining, which reveals the distinctive morphology of Corynebacterium together with additional determination of the bacterium’s physiological properties, such as growth in specific culture media, colony size and shape, pigmentation, cell motility, temperature, and pH. Further analyses include biochemical and enzymatic reactions, such as those determined in automated systems for rapid species identification. The three most frequently used systems are: (i) API Coryne (version 2.0; bioMérieux, France), (ii) BIOLOG (Biolog, Inc., Hayward, CA), and (iii)  RapID CB Plus (Remel, Inc., Norcross, GA). Most of the biochemically based phenotypic analyses are carried out with the test included in the API Coryne system. A brief description of the test’s reactions and corresponding protocol is provided in Section 6.2.2.2, and a description of the protocol for the BIOLOG system in Section 6.2.2.1. The performances of these systems have been evaluated in different studies with variable results because in most cases additional biochemical assays specific for corynebacteria, such as DNase activity and tyrosine hydrolysis, are necessary for definitive identification of the isolates in question.47–49 Most of the chemotaxonomic analyses exploit the presence in corynebacteria of the isomeric form of diaminopimelic acid50 or the bacterial content of whole-cell sugars51 or lipids. Alternatively, they are based on the bacteria’s fatty acids profile or the presence of corynomycolic acids and other β-hydroxy-α-branched carboxylic acids.52 6.1.2.2  Protein Analysis Multilocus Enzyme Electrophoresis (MEE). This technique allows the detection of amino acid substitutions in cellular enzymes that affect their conformation and overall charge. Mobility variants (electromorphs) of the same enzyme can be visualized by their activity in a gel (i.e., as bands of altered migration rates), and each electromorph

Lipo L

L NL NL L

NL NL NL

NL

NL NL NL NL

L NL L NL

Descriptions

105 CFU of A. urinae, the only isolate, per mL. His urinary symptoms resolved and his mental status improved by hospital day 3.26 In another case, a 58-year-old person suffered from several days of dysuria, increased urinary frequency, and nocturia. Urinalysis revealed no bacteria, trace amounts of leukocyte

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Aerococcus

esterase, and negative results for nitrites, protein, ketones, bilirubin, and blood. A second urinalysis demonstrated a moderate level of leukocyte esterase, and negative urine chemistries. The urine culture on the second urine sample later grew >105 CFU of A. urinae, the only isolate, per mL.26 The case studies indicate that A. urinae is a rarely reported pathogen, possibly due to difficulties in the identification of the organism.26 Several conventional and molecular methods are therefore used for detection of the pathogen. In a case of urinary tract infections caused by A. urinae, the patient was treated empirically with intravenous ciprofloxacin.26 Urine cultures grew >105 CFU of A. urinae, the only isolate, per mL. His urinary symptoms resolved and his mental status improved by hospital day 3. The patient was discharged after oral ciprofloxacin was prescribed. Results of repeated urinalysis after discharge were normal. A. viridans strains are naturally susceptible to penicillin, macrolides and related drugs, tetracyclines, and chloramphenicol, and they are intrinsically resistant at a low level to aminoglycosides.1 In a case of endocarditis caused by A.  viridans in a 10-year-old child, he received ampicillin (200 mg/kg/day) and gentamicin (7.5 mg/kg/day) besides decongestive measures. Since the patient’s condition deteriorated during hospital stay, amikacin (15 mg/kg/day) and norfloxacin (15 mg/kg/day) were started on sixth day based on antibiogram. Improvement was noticed from 11th day of hospital stay with a gradual return of fever to baseline. However, at this point the patient developed severe chest pain and a pericardial rub. Echocardiography confirmed the presence of a pericardial effusion. The pericardial fluid was tapped. It was found to be sterile on culture, and steroids were started. The effusion gradually disappeared over the next 5 days, and the cardiac failure was also controlled. Amikacin and norfloxacin were continued for a total duration of 3 weeks.27 These case studies indicate that the treatment is often case sensitive and also depends on the species and its antibiotic susceptibility. A. viridans was found to be resistant to at least one antibiotic amongst eight wild type strains: erythromycin (six strains), tetracycline and minocycline (five strains), chloramphenicol (one strain), and high levels of streptomycin (one strain).1 Augustine et al.27 reported a case of endocarditis caused by A. viridans with multidrug resistance, that is, resistance to penicillin, ampicillin, cefotaxime, gentamicin, and intermediate resistant to ciprofloxacin, but they did not discuss minimum inhibitory concentrations (MICs). According to the antimicrobial susceptibility data of 30 aerococcal isolates obtained from Centers for Disease Control and Prevention,14 the MICs for nine strains were 0.5 g or more of penicillin per mL, with MICs for five strains being more than 1 g/mL; therefore, approximately 46% of aerococci tested were either relatively resistant or resistant to penicillin. Moreover, Christensen et al.28 recently documented that penicillin resistance should be the peculiar characteristic of A. viridans capable of differentiating it from Aerococcus-like organisms.19 Certainly, further investigations are needed to establish the optimal treatment for this pathogen.

In a case of endocarditis caused by A. urinae, treatment modalities were available for 10/11 patients.25 Among nine of these patients treated with a combination of a beta-lactam and an aminoglycoside, seven died. The remaining patient was treated with a beta-lactam alone and survived with neurologic sequelae. In all lethal cases, death occurred during the first 4 weeks (mean, 12 days). It is suggested that all patients are treated intravenously for a minimum of 6 weeks, since all surviving patients, including the presented patient, were treated for this time period either with a beta-lactam alone or with a beta-lactam and an aminoglycoside.

20.1.3  Diagnosis The concept of polyphasic taxonomy integrates different kinds of data and information on microorganisms, that is, phenotypic, genotypic, and phylogenetic, and this method has gained increasing acceptance since its introduction.29 This method includes transmission electron microscopy, fatty acid methyl ester analysis, multilocus enzyme electrophoresis, antimicrobial susceptibility tests, and comparison of 16S rRNA and/or other housekeeping gene(s) sequence data, and ribotyping. After the discovery of A. viridans, all the other strains were discovered on the basis of polyphasic taxonomy. Colony Morphology, Staining, and Biochemical Tests. The presumptive identification of aerobic gram-positive alpha-hemolytic cocci and the decision on whether to more fully identify the isolates are frequently based on Gram stain, colony appearance, and catalase reaction. Because A. urinae is catalase-negative, it could be mistaken for alpha-hemolytic streptococci or enterococci, which are more common urine isolates. At 24 h, the colony morphology resembles that of an alpha-hemolytic streptococcus or lactobacillus; at 48 h, it is similar to that of an Enterococcus. The Gram stain should be differential because A. urinae forms pairs, tetrads, and clusters. However, since A. urinae shows smaller cocci and fewer tetrads than A. viridans does, it could be confused with pediococci or densely packed streptococci or enterococci. The most important routine tests are detection of leucine arylamidase, β-glucuronidase, PYR, hydrolysis of hippurate, and antibiotic susceptibility patterns.7,11,18,26,30,31 Rapid PYR testing is useful for distinguishing between A. viridans or enterococci (both PYR positive) and A. urinae (PYR negative). A Gram stain should be carefully examined for the characteristic arrangement in clusters and tetrads to rule out lactobacillus and other streptococcus. Pediococci are PYR negative and have a Gram stain morphology similar to that of A. urinae; however, they differ in their resistance to vancomycin and in their positive bile esculin test result.18,26 Transmission Electron Microscopy. The transmission electron microscope (TEM) operates on the same basic principles as the light microscope but uses electrons as “light source.” Their much lower wavelength makes it possible to get a resolution a thousand times better than with a light microscope. Cells of both A. urinae and A. viridans had a mean diameter of 0.90–1.10 μm, and strains of both species

214

showed a surface layer.32 In contrast to A. viridans, about 30% of all A. urinae cells showed signs of asymmetrical cell division.32 Sample preparation for TEM requires cultures in log phase. The cells can be fixed for 2 h in 3% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.2, and then pelleted (8800 × g for 2 min). The precipitates should be enrobed in melted Noble agar (Difco) and small cubes with visible clusters of cells can be transferred to 3% glutaraldehyde and fixed overnight at 4°C. The specimens may be post fixed for 90 min in 1% (w/v) OsO4 in cacodylate buffer and stained with 2% (w/v) uranyl acetate in barbiturate buffer. The procedures for the dehydration, embedding in Epon (glycide ether) and the further preparation of thin sections can be performed as described by Blom et al.33 The stained and loaded samples may be viewed using TEM. Antimicrobial Susceptibility Tests. For this, the antimicrobial contained in a reservoir is allowed to diffuse out into the medium (e.g., blood agar) and interact in a plate freshly seeded with the test organisms. A variety of antimicrobial containing reservoirs are used, but the antimicrobial impregnated absorbent paper disc is by far the commonest type used (disc diffusion method). The results of in vitro antibiotic susceptibility testing guide clinicians in the appropriate selection of initial drugs used for individual patients in specific situations. The selection of an antibiotic panel for susceptibility testing is based on the commonly observed susceptibility patterns and is revised periodically. According to the antimicrobial susceptibility data of 30 aerococcal isolates obtained from Centers for Disease Control and Prevention,14 the MICs for nine strains were 0.5 g or more of penicillin per mL, with MICs for five strains being more than 1 g/mL; therefore, approximately 46% of aerococci tested were either relatively resistant or resistant to penicillin. Moreover, Christensen et al.19,28 documented that penicillin resistance should be the peculiar characteristic of A. viridans, capable of differentiating it from Aerococcus-like organisms. Table 20.1 shows the antibiotic susceptibilities of A. urinae, A. viridans, and A. sanguinicola. Fatty Acid Methyl Ester Analysis. Microorganisms can be classified based on gas chromatographic analysis of extracted microbial fatty acid methyl esters (FAMEs). Microbial fatty acid profiles are unique from one species to another, and this has allowed for the creation of very large microbial libraries. Location of the double-bond position of monounsaturated fatty acids of A. viridans was accomplished by combined gas chromatography (GC)–mass spectrometry analysis of dimethyl disulfide (DMDS) derivatives. The monoenoic fatty acids from whole bacterial cells were converted to methyl esters and then to DMDS adducts and analyzed by capillary GC–mass spectrometry.37,38 Multilocus Enzyme Electrophoresis (MLEE). Isolates are characterized by the relative electrophoretic mobilities of a large number of water-soluble cellular enzymes. Because the net electrostatic charge and, hence, the rate of migration of a protein during electrophoresis are determined by its amino acid sequence, mobility variants (electromorphs or

Molecular Detection of Human Bacterial Pathogens

TABLE 20.1 Antibiotic Susceptibilities of A. urinae and A. viridans. Data Has Been Adapted from Observations Reported Earliera Antibiotic Penicillin G Trimethoprimsulfamethoxazole Vancomycin Gentamicin Tetracycline Ciprofloxacin Ampicillin Erythromycin Cefuroxime Chloramphenicol

A. urinae

A. viridans

A. sanguinicola

S R

R S

S NA

S R S S S R S S

S S/R S R R R R R

S NA NA NA S S S NA

Abbreviations:  S, susceptible; R, resistant; NA, not available. Source: Uh, Y. et al., J. Korean Med. Sci., 17, 113, 2002; Zhang, Q. et al., J.  Clin. Microbiol., 38, 1703, 2000; Skov, R.L. et al., Diagn. Microbiol. Infect. Dis., 21, 219, 1995; Ibler, K. et al., Scand. J. Infect. Dis., 40, 761, 2008; Colakoglu, S. et al., Infection, 36, 288, 2008.

allozymes) of an enzyme can be directly equated with alleles at the corresponding structural gene locus.39 Christensen et al.32 used various enzymes for generating MLEE patterns and finally constructed UPGMA and Minimal Evolution trees based on the results. They found that the tree concurs with the ribotyping data and distinguished A. urinae strains as a group of strains distinct from the type strain of A. viridans. Sample preparation methods for MLEE have been adapted from those described by Selander et al.39 For this technique, whether cells are grown in broth or on agar plates is unimportant, since growth conditions do not affect the electrophoretic mobilities of enzymes. However, some enzymes may vary in activity (but not in mobility) on gels, depending on the type of medium in which the cells were grown. The activity level of certain enzymes may also vary with the composition of the culture medium, but, again, electrophoretic mobilities are not affected. Cells are normally lysed by sonication; however, any method of lysis that does not denature proteins may be used. After lysis and centrifugation at 30,000 × g for 20 min, aliquots of the several milliliters of lysate (supernatant) are transferred to three or four culture tubes and stored at −70°C until used for electrophoresis. At that temperature, lysates can be stored for several years without significant loss of activity of most enzymes. The stability of the enzymes varies markedly among species of bacteria, and so this step may be avoided. Apparatus for horizontal starch-gel electrophoresis is used. Starch gels are preferred over polyacrylamide gels. To prepare a gel, a suspension of starch and gel buffer is heated just beyond the boiling point.39 The suspension is immediately poured into a lucite gel mold. After the gel has cooled at room temperature for 2 h, it is wrapped in plastic film to prevent desiccation. Gels are used within 24 h of preparation. In loading a gel, pieces of Whatman no. 3 filter

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paper (9 mm by 6 mm) are individually dipped into samples of lysate, blotted on filter paper to remove excess liquid, and then inserted at 3-mm intervals in a continuous slit cut in the gel. Up to 20 lysates can be electrophoresed on a single gel. Pieces of filter paper dipped in amaranth dye are inserted at one or both ends of the slit to mark the migration front of the buffer line. During electrophoresis, a constant voltage is maintained, and the gel is cooled by a pan of ice supported above the gel mold on a thin plate of glass. Gels may be run in cold rooms or refrigerators at 4°C. Following electrophoresis, three or four horizontal slices (1–2 mm thick) are cut from the gel with a thin wire and incubated individually at 37°C in various selective enzyme staining solutions.39 The activity level and degree of separation of electromorphs of an enzyme may depend on the type and pH of the buffer system used and, to some extent, on the concentration of starch in the gel. Thus, an enzyme may appear to be monomorphic or polymorphic for only two or three electromorphs in one buffer system but exhibit 5–10 electromorphs in another buffer system. Occasionally, the relative mobilities of certain electromorphs are reversed in different buffer systems. Comparisons of the mobilities of enzymes from different isolates are made visually against one another on the same gel slice. It is not sufficient to compare relative mobilities of an enzyme in different strains by measuring distances of migration from the origin, even when the enzymes have been electrophoresed on the same gel. Each isolate is characterized by its combination of electromorphs over the number of enzymes assayed, and distinctive profiles of electromorphs, corresponding to unique multilocus genotypes, are designated electrophoretic types (ETs), which are equivalent to allele profiles. For analysis, a distance matrix in MEGA format may be generated using the computer program ETMEGA of T.S. Whittam (www.foodsafe.msu.edu/whittam/#programs) as mentioned by Chritensen et al.32 ETMEGA calculates the genetic distance between pairs of “electrophoretic types,” which were defined by distinct patterns of electrophoretic mobilities of the 11 housekeeping enzymes. Distance is measured as the proportion of mismatched loci between pairs of ETs. Null alleles are not used in the calculation of pair wise distances. Phylogenetic and molecular evolutionary analyses using this distance matrix may be conducted using MEGA version 4.32,40–42 Ribotyping. Ribotyping is a method that can identify and classify bacteria based upon differences in rRNA. It generates a highly reproducible and precise fingerprint that can be used to classify bacteria from the genus through and beyond the species level. DNA is extracted from a colony of bacteria and then restricted into discrete-sized fragments. In experiments performed by Christensen et al.32 ribotyping of the 26 A. urinae strains and the type strain of A. viridans with EcoR1 and HaeIII resulted in large DNA fragments of more than 40 kb, which could not be resolved using the RiboPrinter system. Ribotyping these strains with XbaI resulted in seven distinct ribotype patterns, each comprising between 1 and 12 strains. They showed that strains of A. urinae were clearly distinct from A. viridansT.

20.2  METHODS 20.2.1  Sample Preparation For extraction of Aerococcus DNA, a Wizard SV genomic DNA purification system (Promega, Madison, Wis.) may be used with some modifications. The bacterial pellet is resuspended in 400 μl enzymatic lysis solution (47 mM EDTA, 25 mg/ml lysozyme [Sigma, St. Louis, Mo.], 20 μg/ml lysostaphin [Sigma]) and incubated at 37°C for 2 h. Then, proteinase K (19.2 mg/ml; Roche) is added to a final concentration of 0.4 mg/ml and the mixture is incubated at 55°C for 1 h. Nuclei Lysis solution (Promega) and RNase solution (Promega) are added, and after mixing, the reaction solution is incubated at 80°C for 10 min. Further purification steps are done according to the instructions of the manufacturer. The final elution volume is 200 μl. For each PCR, 2 μl of this DNA extract is applied.

20.2.2  Detection Procedures (i) Aerococcus Viridans-Specific PCR  Martín et al. utilized A. viridans-specific primers AC2 (5′-GTGC TTGCACTTCTGACGTTAGC-3′) and AC4 (5′-TGAGC CGTGGGCTTTCACAT-3′) to amplify a 540 bp fragment from 16S rRNA gene.43 Furthermore, Grant et al. developed a rapid protocol for identification of A. viridans through PCR amplification of a 16S rRNA gene fragment, followed by detection with an oligonucleotide probe.44 (ii) PCR and Sequencing  Sequencing analysis of 16S rRNA gene allows identification of all Aerococcus species.45 In this protocol, a large fragment of the 16S rRNA gene (corresponding to positions 30 ± 1521 of the Escherichia coli 16S rRNA gene) is amplified by PCR using conserved primers close to the 3′ and 5′ ends of the gene. The PCR product is directly sequenced using a Taq Dye-Deoxy Terminator Cycle Sequencing kit (Applied Biosystems) and an automatic DNA sequencer. (iii) PCR Ribotyoing  PCR ribotyping was developed by Kostman et al.46 and Gurtler et al.47 in the 1990s as a response in part to the need in the clinical microbiology laboratory setting for expeditious epidemiological discrimination among pathogenic microorganisms without the use of probes, thus making the analysis more widely applicable.48 Primers complementary to the 3′ end of the 16S rRNA gene and the 5′ end of the 23S rRNA gene were used such that the amplified fragment revealed length heterogeneity of the internal spacer regions (ISR) located between the 16S and 23S genes of the six rRNA gene operons.49 However, PCR ribotyping proved not to be universally applicable.50 In some species, ISR length is variable within and between isolates, in others ISR lengths are limited to one or two sizes, usually dependent on the number of tRNAs present.48 In an attempt to further subtype isolates for which PCR ribotyping was nondiscriminatory, Ryley et al.51 and Shreve et al.52 performed PCR amplification of ISRs, as described above, followed by restriction endonuclease

216

Molecular Detection of Human Bacterial Pathogens

Tissue or Cells

Quantitation by Nanodrop

Genomic DNA Extraction Genomic DNA Quantitation by Electrophoresis

Restriction digestion and size separation of DNA fragments

Hybridization with RNA probe

Dendrogram showing similarities between ribotypes

FIGURE 20.1  Demonstration of ribotyping. DNA is isolated from a cultured isolate, restricted and size separated in an agarose gel. The gel is then hybridized with labeled rRNA probe from a type strain of the species or genus, which binds to fragments containing copies of the rRNA operon. Following probe detection, fragments with bound probe are visualized, forming a characteristic RFLP. The advantages of ribotyping over other RFLP methods is that both highly variable and conserved sequences are highlighted by the probe, which further makes the method suitable to type strains over a broad phylogenetic range, and the same process can be used for analysis of strains within any species.

digestion. Christensen et al.32 found the following enzymes suitable for Aerococcus: XbaI, EcoR1, and HaeIII. An alternate variation of PCR ribotyping, amplified rRNA gene restriction analysis (ARDRA) is based on PCR amplification of the 16S rRNA gene followed by restriction digestion. Jayarao et al.53 developed this technique to determine subspecies of Streptococcus uberis, as a means of avoiding methods involving DNA hybridization or sequencing. Later, as described by Dupont Qualicon, the RiboPrinter automated ribotyping system came as a technological breakthrough with respect to convenience, reproducibility, and speed. The speed in this case appears to be two- to threefold greater than that for manual ribotyping, with far less labor-intensive procedures. For polyphasic characterization of A. urinae, automated ribotyping method (Figure  20.1) was used successfully with the Riboprinter (Qualicon Inc., Wilmington, DE).32

two distinct lineages of A. urinae exist, which can be identified by phenotypic markers. Future recognition of these biovars may reveal potential differences in ecology and clinical significance. In daily routine practice, species identification of Aerococcus causes problems. Routine diagnostic procedures should be able to identify this bacteria, and clinicians should be aware of the pathogenic potential of this organism.

ACKNOWLEDGMENTS We thank Dr. Mark Lawrence and Dr. Shane Burgess for their support in writing the manuscript. We are also grateful to Dr. Amit Ghosh, and Dr. Walter J. Diehl for encouragement.

REFERENCES

20.3  CONCLUSION The genus Aerococcus shows distinct interspecies differences, but constitutes a well-circumscribed genus as revealed by 16S rRNA sequence and other data. Phenotypic and molecular analyses demonstrate that A. urinae is to some extent a heterogeneous species belonging to the genus Aerococcus. At least



1. Buu-Hoi, A., Le Bouguenec, C., and Horaud, T., Genetic basis of antibiotic resistance in Aerococcus viridans, Antimicrob. Agents Chemother., 33, 529, 1989. 2. Astudillo, L. et al., Spondylodiscitis due to Aerococcus urinae: A first report, Scand. J. Infect. Dis., 35, 890, 2003. 3. Santos, R. et al., Lymphadenitis caused by Aerococcus urinae infection, Scand. J. Infect. Dis., 35, 353,2003. 4. Williams, R.E., Hirch, A., and Cowan, S.T., Aerococcus, a new bacterial genus, J. Gen. Microbiol., 8, 475, 1953.

Aerococcus







5. Lawson, P.A., Falsen, E., Ohlen, M., and Collins, M.D., Aerococcus urinaehominis sp. nov., isolated from human urine, Int. J. Syst. Evol. Microbiol., 51, 683, 2001. 6. Aguirre, M., and Collins, M.D., Phylogenetic analysis of some Aerococcus-like organisms from urinary tract infections: Description of Aerococcus urinae sp. nov., J. Gen. Microbiol., 138, 401, 1992. 7. Christensen, J.J. et al., Bacteremia/septicemia due to Aerococcus-like organisms: Report of seventeen cases. Danish ALO Study Group, Clin. Infect. Dis., 21, 943, 1995. 8. Christensen, J.J., Korner, B., and Kjaergaard, H., Aerococcuslike organism—an unnoticed urinary tract pathogen, Apmis, 97, 539, 1989. 9. Sierra-Hoffman, M., Watkins, K., Jinadatha, C., Fader, R., and Carpenter, J.L., Clinical significance of Aerococcus urinae: A retrospective review, Diagn. Microbiol. Infect. Dis., 53, 289, 2005. 10. Lawson, P.A., Falsen, E., Truberg-Jensen, K., and Collins, M.D., Aerococcus sanguicola sp. nov., isolated from a human clinical source, 51, 475, 2001. 11. Collins, M.D., Jovita, M.R., Hutson, R.A., Ohlen, M., and Falsen, E., Aerococcus christensenii sp. nov., from the human vagina, Int. J. Syst. Bacteriol., 49 Pt 3, 1125, 1999. 12. Collins, M.D., Jovita, M.R., Hutson, R.A., Ohlen, M., and Falsen, E., Note: Aerococcus christensenii sp. nov., from the human vagina, Int. J. Syst. Bacteriol., 49, 1125, 1999. 13. Christensen, J.J. et al., Aerococcus urinae: Intraspecies genetic and phenotypic relatedness, Int. J. Syst. Bacteriol., 47, 28, 1997. 14. Bosley, G.S. et al., Phenotypic characterization, cellular fatty acid composition, and DNA relatedness of aerococci and comparison to related genera, J. Clin. Microbiol., 28, 416, 1990. 15. Schuur, P.M.H. et al., Urinary tract infections with Aerococcus urinae in the south of The Netherlands, J. Urol., 161, 1414, 1999. 16. Martin, V. et al., Characterization of Aerococcus viridans isolates from swine clinical specimens, J. Clin. Microbiol., 45, 3053, 2007. 17. Razeq, J.H., Thomas, G.M., and Alexander, D., The first reported case of Aerococcus bacteremia in a patient with HIV infection, Emerg. Infect. Dis., 5, 838, 1999. 18. Facklam, R., and Elliott, J.A., Identification, classification, and clinical relevance of catalase-negative, gram-positive cocci, excluding the streptococci and enterococci, Clin. Microbiol. Rev., 8, 479, 1995. 19. Uh, Y., Son, J.S., Jang, I.H., Yoon, K.J., and Hong, S.K., Penicillin-resistant Aerococcus viridans bacteremia associated with granulocytopenia, J. Korean Med. Sci., 17, 113, 2002. 20. Kern, W., and Vanek, E., Aerococcus bacteremia associated with granulocytopenia, Eur. J. Clin. Microbiol., 6, 670, 1987. 21. Swanson, H., Cutts, E., and Lepow, M., Penicillin-resistant Aerococcus viridans bacteremia in a child receiving prophylaxis for sickle-cell disease, Clin. Infect. Dis., 22, 387, 1996. 22. Hermansson, K. et al., Structural studies of the capsular polysaccharide from Aerococcus viridans var. homari, Carbohydr. Res., 208, 145, 1990. 23. Nasoodi, A., Ali, A.G., Gray, W.J., and Hedderwick, S.A., Spondylodiscitis due to Aerococcus viridans, J. Med. Microbiol., 57, 532, 2008. 24. Tekin, A., Tekin, G., Turunc, T., Demiroglu, Z., and Kizilkilic, O., Infective endocarditis and spondylodiscitis in a patient due to Aerococcus urinae: first report, Int. J. Cardiol., 115, 402, 2007. 25. Ebnother, C., Altwegg, M., Gottschalk, J., Seebach, J.D., and Kronenberg, A., Aerococcus urinae endocarditis: Case report and review of the literature, Infection, 30, 310, 2002.

217 26. Zhang, Q., Kwoh, C., Attorri, S., and Clarridge, J.E., Aerococcus urinae in urinary tract infections, J. Clin. Microbiol., 38, 1703, 2000. 27. Augustine, T., Thirunavukkarasu, Bhat, B.V., and Bhatia, B.D., Aerococcus viridans endocarditis. Case report, Indian Pediatr., 31, 599, 1994. 28. Christensen, J.J., Korner, B., Casals, J.B., and Pringler, N., Aerococcus-like organisms: Use of antibiograms for diagnostic and taxonomic purposes, J. Antimicrob. Chemother., 38, 253, 1996. 29. Vandamme, P. et al., Polyphasic taxonomy, a consensus approach to bacterial systematics, Microbiol. Rev., 60, 407, 1996. 30. Christensen, J.J., Vibits, H., Ursing, J., and Korner, B., Aerococcus-like organism, a newly recognized potential urinary tract pathogen, J. Clin. Microbiol., 29, 1049, 1991. 31. Kristensen, B., and Nielsen, G., Endocarditis caused by Aerococcus urinae, a newly recognized pathogen, Eur. J. Clin. Microbiol. Infect. Dis., 14, 49, 1995. 32. Christensen, J.J. et al., Aerococcus urinae: Polyphasic characterization of the species, Apmis, 113, 517, 2005. 33. Blom, J., Mansa, B., and Wilk, A., A study of Russell bodies in human monoclonal plasma cells by means of immunofluorescence and electron microscopy, Acta Pathol. Microbiol. Scand., A 84, 335, 1976. 34. Skov, R.L., Klarlund, M., and Thorsen, S., Fatal endocarditis due to Aerococcus urinae, Diagn. Microbiol. Infect. Dis., 21, 219, 1995. 35. Ibler, K. et al., Six cases of Aerococcus sanguinicola infection: Clinical relevance and bacterial identification, Scand. J. Infect. Dis., 40, 761, 2008. 36. Colakoglu, S. et al., Three cases of serious infection caused by Aerococcus urinae: A patient with spontaneous bacterial peritonitis and two patients with bacteremia, Infection, 36, 288, 2008. 37. Moss, C.W., and Daneshvar, M.I., Identification of some uncommon monounsaturated fatty acids of bacteria, J. Clin. Microbiol., 30, 2511, 1992. 38. Moss, C.W., Wallance, P.L., and Bosley, G.S., Identification of monounsaturated fatty acids of Aerococcus viridans with dimethyl disulfide derivatives and combined gas chromatography-mass spectrometry, J. Clin. Microbiol., 27, 2130, 1989. 39. Selander, R.K. et al., Methods of multilocus enzyme electrophoresis for bacterial population genetics and systematics, Appl. Environ. Microbiol., 51, 873, 1986. 40. Kumar, S., Nei, M., Dudley, J., and Tamura, K., MEGA: A biologist-centric software for evolutionary analysis of DNA and protein sequences, Brief Bioinform., 9, 299, 2008. 41. Kumar, S., Tamura, K., and Nei, M., MEGA3: Integrated software for Molecular Evolutionary Genetics Analysis and sequence alignment, Brief Bioinform., 5, 150, 2004. 42. Tamura, K., Dudley, J., Nei, M., and Kumar, S., MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0, Mol. Biol. Evol., 24, 1596, 2007. 43. Martín, V. et al., Characterization of Aerococcus viridans isolates from swine clinical specimens, J. Clin. Microbiol., 45, 3053, 2007. 44. Grant, K.A. et al., Rapid identification of Aerococcus viridans using the polymerase chain reaction and an oligonucleotide probe, FEMS Microbiol. Lett., 95, 63, 2006. 45. Lawson, P.A. et al., Aerococcus urinaehominis sp. nov., isolated from human urine, Int. J. Syst. Evol. Microbiol., 51, 683, 2001.

218 46. Kostman, J.R. et al., A universal approach to bacterial molecular epidemiology by polymerase chain reaction ribotyping, J. Infect. Dis., 171, 204, 1995. 47. Gurtler, V., and Stanisich, V.A., New approaches to typing and identification of bacteria using the 16S-23S rDNA spacer region, Microbiology, 142, 3, 1996. 48. Bouchet, V., Huot, H., and Goldstein, R., Molecular genetic basis of ribotyping, Clin. Microbiol. Rev., 21, 262, 2008. 49. Gurtler, V., and Barrie, H.D., Typing of Staphylococcus aureus strains by PCR-amplification of variable-length 16S-23S rDNA spacer regions: characterization of spacer sequences, Microbiology, 141, 1255, 1995. 50. Cartwright, C.P., Polymerase chain reaction ribotyping: A “universal” approach? J. Infect. Dis., 172, 1638, 1995.

Molecular Detection of Human Bacterial Pathogens 51. Ryley, H.C., Millar-Jones, L., Paull, A., and Weeks, J., Characterisation of Burkholderia cepacia from cystic fibrosis patients living in Wales by PCR ribotyping, J. Med. Microbiol., 43, 436, 1995. 52. Shreve, M.R., Johnson, S.J., Milla, C.E., Wielinski, C.L., and Regelmann, W.E., PCR ribotyping and endonuclease subtyping in the epidemiology of Burkholderia cepacia infection, Am. J. Respir. Crit. Care. Med., 155, 984, 1997. 53. Jayarao, B.M., Dore, J.J., Jr.. and Oliver, S.P., Restriction fragment length polymorphism analysis of 16S ribosomal DNA of Streptococcus and Enterococcus species of bovine origin, J. Clin. Microbiol., 30, 2235, 1992.

21 Bacillus Noura Raddadi, Ameur Cherif, and Daniele Daffonchio CONTENTS 21.1 Introduction.......................................................................................................................................................................219 21.1.1 Classification, Morphology, and Biology of Bacillus spp. ...................................................................................219 21.1.2 Genetic Baseline of B. cereus Group Ecotypes.................................................................................................... 220 21.1.3 Species Discrimination in B. cereus Group......................................................................................................... 220 21.1.4 Clinical Features and Toxicology of Human Pathogenic Bacillus spp................................................................ 220 21.1.4.1 Bacillus anthracis.................................................................................................................................. 220 21.1.4.2 Bacillus cereus and Other Human Pathogenic Bacillus spp. ............................................................... 221 21.1.5 Diagnosis Techniques........................................................................................................................................... 223 21.1.5.1 Conventional Diagnosis Techniques...................................................................................................... 223 21.1.5.2 Molecular Diagnosis Techniques........................................................................................................... 223 21.2 Methods............................................................................................................................................................................ 224 21.2.1 Sample Preparation............................................................................................................................................... 224 21.2.2 Detection Procedures............................................................................................................................................ 224 21.3 Conclusions and Future Perspectives............................................................................................................................... 225 References.................................................................................................................................................................................. 226

21.1  INTRODUCTION 21.1.1  C  lassification, Morphology, and Biology of Bacillus spp. Bacteria in the genus Bacillus are gram-positive, spore formers, and rod-shaped. They are very diverse in terms of physiology, ecological niche, genes sequences, and regulation. Their impact on human activity varies from probiotic effect1,2 to severe pathogenecity. The oldest, most infectious, and potentially lethal human disease is the anthrax that is caused by Bacillus anthracis.3 However, many other species within this genus have emerged as new human pathogens associated with foodborne diseases that can cause severe and even fatal infections. These include B. cereus, B. weihenstephanenesis, B. pumilus, B. mojavensis, B. licheniformis, B. subtilis, and B. circulans. All Bacillus species, due to endospore formation, can survive heat treatment and disinfection procedures and hence pose serious health concerns as contaminants of food or in hospital settings as a cause of nosocomial infections,4–10 as well as severe illnesses, especially in immunocompromised patients or immunocompetent individuals with risk factors such as intravenous drug use, hemodialysis, and leukemia. The production of toxins and enzymes by these bacteria is considered the causative agent of several human diseases. In this chapter, we survey the different human pathologies caused by Bacillus spp., mainly those of the B. cereus group.

Before giving details on the clinical features, toxicology, and the methods of detection of these bacteria and their corresponding pathogenecity factors, we will briefly introduce the complex ecology and the identification challenges that characterize several Bacillus species. The B. cereus group, which shows a “bivalent face” in terms of its important impact on human activity will be used as a paradigm.11,12 B. anthracis is the most serious human pathogen in this group and the major subject of this chapter. This bacterium is the etiological agent of anthrax, a potentially fatal disease for humans and animals, and a bioterrorism agent. B. cereus is the most known foodborne pathogen that has been associated with food-poisoning illnesses13,14 and other kinds of clinical infections that are reported in this chapter. 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. weihenstephanensis, a psychrotolerant species capable of growing at temperatures as low as 4°C–6°C is implicated in food spoilage.15 In addition to B. anthracis, B. cereus, B. thuringiensis, and B. weihenstaephanenesis, the B. cereus group encompasses two other species, B. mycoides and B. pseudomycoides, which 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 studies16–18 and the sequencing of the ribosomal RNA 219

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genes.19–21 However, a marked variability is always observed when large collections of strains are examined by DNA fingerprinting methods that target the whole genome22–26 and/or discrete genes.27–31 Hence, the phylogenetic and taxonomic relationships among these species are still open to debate. The genetic variability observed within and between these species has raised several interesting questions and poses 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?

21.1.2  Genetic Baseline of B. cereus Group Ecotypes The ecology of these bacteria is still far from clear, despite several recent analyses that have shown that a major environmental niche for these bacteria is the invertebrate intestinal tract.32–34 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. thuringiensis, and in mammals and humans for B. anthracis and B. cereus.35 Apart from this general scheme, the ecology of these species outside the invertebrate host is not completely clear, and the ability of several environments with which these bacteria have been commonly associated, to continuously support their life cycle has been questioned. It has been shown that with respect to other soil-inhabiting bacteria, such as 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, such as animal guts or animal tissues and fluids, rather than the plant environment.36,37 Specialization on different “animal” niches has lead to different evolution of the species in the 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 pathways38: 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. 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 for lethal virulence.

21.1.3  Species Discrimination in B. cereus Group The similarity among such closely related bacteria as the species of the B. cereus group posed serious challenges for species discrimination. This is an important point to address, considering the differences in the virulence potential of these microorganisms. Typically, B. anthracis can be

Molecular Detection of Human Bacterial Pathogens

differentiated from most of B. cereus and B. thuringiensis strains by specific tests, such as penicillin G and gamma phage sensibility, lack of motility, and beta-hemolysis on blood agar. However, 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.39 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.39 Thus, considering the virulence potential of strains that are genetically the near neighbors of B. anthracis, approaches that might rapidly identify these strains are of great interest. From a safety perspective, these strains could represent alternative hosts for B. anthracis toxin genes.39 Several B. cereus strains resulted, strictly related to B. anthracis. For example, Keim et al.38 and Radnedge et al.40 individuated by amplified fragment-length polymorphism (AFLP) B. cereus and B. thuringiensis strains closely related to B. anthracis. Radnedge et al.41 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 for the identification of species borderline strains, like, among others, rep-PCR12 and multilocus sequence typing (MLST)42 or, recently, comparative genomics43 and microarray analysis.44 Alternative approaches have been based on length or sequence polymorphisms in variable‑number tandem repeats in multiple loci (multilocus VNTR analysis [MLVA])45 or on signature single nucleotide polymorphisms (SNPs) in the genome.46 MLVA has been used as a gold standard for subtyping B. anthracis isolated worldwide (see among others35,45,47,48). Together with MLVA, SNP analysis is greatly contributing in typing and tracing B. anthracis isolates, especially by the whole-genome SNP analysis.49 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.50 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. Gierczyn´ski et al.51 developed a simple and cost-effective restriction-siteinsertion PCR (RSI-PCR)–based assay that precisely detects the B. anthracis-specific nonsense mutation in plcR gene by restriction digestion with the endonuclease SspI.

21.1.4  C  linical Features and Toxicology of Human Pathogenic Bacillus spp. 21.1.4.1  Bacillus anthracis B. anthracis, the causative agent of anthrax disease, has been well studied for over 150 years and continues to be a threat as a biological weapon.3 This bacterium, forms spores (1–2

Bacillus

μm in size) able to withstand several kinds of stresses that are usually lethal to vegetative cells, such as ultraviolet and ionizing radiation, heat, various chemicals,52 and oxidative stress.53 Anthrax is primarily a disease affecting herbivores, but it can be transmitted to all mammals, including humans. Typically, animals become infected with B. anthracis by ingesting spores while grazing on contaminated soil or feed and also through the skin via biting insects.54 Transmission of the spores to humans occurs through three routes of entry: (i) cutaneously, by direct contact with infected animals or by handling infected animal products55; (ii) respiratory, by inhalation of sufficient quantity of spores; and (iii) oropharyngeal and/or gastrointestinal (GI), by the ingestion of undercooked, contaminated meat.56 All three forms of anthrax can be fatal if not treated in time, but the cutaneous form is often self-limiting.57 A typical anthrax skin lesion corresponds to an ulcer covered by a characteristic black eschar.58 The pathogenecity of B. anthracis depends on two major factors: a poly-γ-d-glutamic acid capsule and the three-component secreted anthrax toxin codified by plasmids pXO2 and pXO1, respectively. In addition to protecting the bacteria from the phagocytosis, the capsule allows bacterial proliferation and subsequent toxemia to develop in the bloodstream because of its poor immunogenic properties. The anthrax toxin is composed of three proteins known as protective antigen (PA), lethal factor (LF), and edema factor (EF). Once in the body, the PA binds to receptors that are present on most cells, including macrophages, and after proteolytic activation, it combines to LF and EF, allowing their delivery inside the cell.59 Transcription of the genes pag, lef, and cya encoding the three-toxin proteins PA, LF, and EF, respectively, is regulated by a gene known as the anthrax toxin activator (atxA); whereas the capsule proteins, codified by the capBCADE operon, are regulated by AcpA and AcpB.60,61 Once in the host and after their phagocytosis by local macrophages, spores germinate and multiply in their way to the regional lymph nodes.41 Along with other mechanisms, two spore-bound superoxide dismutases (SOD15 and SODA1), enzymes that are capable of detoxifying oxygen radicals,62 were recently reported to play a crucial role in protecting the spores against the oxidative burst of macrophages in mouse model.53 Following spore germination and bacterial proliferation, the capacity of the lymph nodes is exceeded; hence, the vegetative bacteria enter the bloodstream, which leads to bacteremia and the distinctive pathology of edema, hemorrhage, and tissue necrosis.59 Dissemination of B. anthracis by hematogenous or lymphatogenous spread from a cutaneous, mediastinal, or GI focus can lead to infection at other sites, such as the brain, allowing entry into the central nervous system (CNS) and development of anthrax meningitis.63 Another form of anthrax meningitis in which no primary focus is found has been reported and is known as “primary anthrax meningitis.”64 With regard to the epidemiology, anthrax infection has been reduced dramatically in developed countries as the result of the widespread use of vaccines in high-risk people

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and animals, as well as the improvements in industrial hygiene. Currently, only very few cases of human anthrax occur naturally in these countries. However, the scenario is different in developing countries such as Africa and Asia, where the disease is endemic; this is due in part to difficulties of implementing vaccination programs. Of the three forms of the disease, cutaneous anthrax is the most frequent, followed by the inhalational and GI or oropharyngeal form.55,57,65,66 Only few case reports have been published regarding anthrax meningitis epidemiology, and most of them report anthrax meningitis in settings of one of the three forms of the disease.57,64,67,68 21.1.4.2  Bacillus cereus and Other Human Pathogenic Bacillus spp. Due to their ubiquitous presence in the environment, Bacillus spp., other than B. anthracis, isolated from clinical specimens have been often considered as a contamination; hence, most laboratories do not identify these isolates further. It has been reported that Bacillus spp. are isolated from 0.1% to 0.9% of blood cultures, but only 5%–10% of these are clinically significant infections.69–71 However, in recent years there has been a growing number of reports dealing with their implication in severe and even lethal infections. Bacillus spp. other than B. anthracis were shown to cause nosocomial bacteremia, pneumonia, brain and liver abscesses, infection of CNS, periodontitis, peritonitis, endophthalmitis, soft-tissue infection, and meningitis. Nosocomial Bacteremia. B. cereus is the most frequently isolated of Bacillus spp. involved in the development of nosocomial infections.7–10,72,73 Van Der Zwet72 described an outbreak of invasive B. cereus infections in a neonatal intensive care unit caused by contaminated balloons used for manual ventilation. These balloons had been cleaned with detergent, which was insufficient to kill spores of B. cereus. Kalpoe et al.9 reported an outbreak of vancomycinresistant B. cereus respiratory tract colonization in pediatric intensive care unit (PICU) patients due to contamination of reusable ventilator air-flow sensors. B. cereus spore contamination of alcohol used to disinfect the air-flow sensors was postulated to be the source of the dissemination. Dohmae et al.8 reported on five bloodstream infections in five patients related to catheter use. B. cereus contamination of reused towels, which would have occurred during the wash process, was considered to be the origin of this outbreak. Ohsaki et al.7 reported on the occurrence of three consecutive cases of B. cereus bacteremia in patients with hematologic malignancies after hospital renovation. The authors considered that filters used in the heating, ventilation, and air-conditioning systems, as well as towels and gowns, were the probable sources of the B. cereus outbreak. Kuroki et al.,10 demonstrated that B. cereus and B.  thuringiensis species previously implicated in nosocomial bacteremia by catheter infection were able to form biofilm after 24 h of incubation. To date, there is one report describing the occurrence of fatal B. thuringiensis bacteremia, in a neutropenic patient who suffered from severe pulmonary disease,74 indicating

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that this entomopathogenic bacterium may behave as an opportunistic pathogen in immunocompromised individuals. B. cereus together with B. licheniformis and B. pumilus were shown to be implicated in 3.4% of 350 bacteremic episodes that occurred during a 5-year period in patients having hemotological malignancies.75 To date, only one case of B. licheniformis nosocomial infection, affecting a very-low-birth-weight neonate in a neonatal intensive care unit, has been reported.76 In immunocompetent individuals, B. licheniformis bacteremia has been reported in association with central venous catheters.77 B. pumilus has also been implicated in persistent bacteremia in a patient with cholangitis and in nosocomial bacteremia via catheter infection.78,79 In addition to antibiotic treatment, the central venous catheter may need to be removed for complete cure. Taken together, these reports indicate that, although cases of Bacillus spp. other than B. anthracis and B. cereus bacteremia are rare in healthy individuals, it is important to consider the potential pathogenicity of Bacillus spp. in immunocompromised hosts, including premature infants and neonates, as well as in patients with various risk factors. Endophthalmitis. Infectious endophthalmitis, the inflammation of intraocular tissues or fluids due to intraocular infection, is a major public health concern. In addition to fungi and gram-negative bacteria80; different Bacillus species, such as B. megaterium,81 B. subtilis, B. mycoides, B. pumilus, B.  flexus, and B. thuringiensis, have been reported as causative agents of infectious endophthalmitis.82 However, the most important eye pathogen in this genus is B. cereus because of its ability to blind rapidly. Endophthalmitis associated to this bacterium often results in significant vision loss or loss of globe architecture in 1–2 days.5,83 The eye pathogenesis of B. cereus is due to rapid growth, migration, and toxin production by this microorganism, which lead to the failure of the host immune defenses.82,84 In addition to toxin production by the offending bacterium, the pathogenesis of B. cereus was also attributed to the intraocular inflammatory response and, in particular, to tumor necrosis factoralpha (TNFα) in experimental endophthalmitis.85 Recently, B. cereus was also shown to induce the permeability of the blood-ocular barrier in experimental endophthalmitis.86 Foodborne Illnesses. Bacillus cereus is well known as a common cause of emetic and diarrheal food poisoning. The symptoms of these foodborne illnesses are usually mild and the disease is often self-limiting. However, this opportunistic human pathogen can be associated with severe intoxications and infections that can be fatal, even in otherwise healthy individuals. Indeed, three fatal cases due to fulminant liver failure following intoxication with cereulide-producing B. cereus strains and liver abscess have been reported to date.13,87,88 Other Bacillus spp. have also been implicated in foodborne illnesses, including B. weihenstephanensis, B. subtilis, B. licheniformis, and B. pumilus; while other species, such as B. mojavensis, B. firmus, B. megaterium, B. simplex, and B. fusiformis were shown to produce toxins with a cereulide-like mode of action.14

Molecular Detection of Human Bacterial Pathogens

Pneumonia. Bacillus cereus has also been associated with fulminant pneumonia in five otherwise healthy metalworkers: a single nonfatal case in 1994 and four fatal cases: two in 1997 and two in 2003.89–91 The B. cereus strains responsible for these cases of fulminant pneumonia were shown to produce a capsule and to have the genetic determinants that codify for the anthrax toxins.92 A sixth case of fatal pneumonia caused in a leukemic patient by carbapenem-resistant B. cereus has also been reported recently.93 The fact that B. cereus has been implicated in such unusual severe fatal pneumonia in immunocompetent individuals highlights the need for increased awareness of the potential for this opportunistic human pathogen to cause serious systemic infections, even in patients who appear to be otherwise healthy. Thus, a better understanding of the environmental and occupational risk factors that may increase the susceptibility of individuals to infection by this bacterium should be considered. Periodontitis. The genus Bacillus has received relatively little attention in oral microbiology, and the Bacillus spp. isolated from human oral samples have been often regarded as transient microflora or as external contaminants. However, there have been several reports on the implication of Bacillus species, such as B. cereus/B. thuringiensis,94 B. pumilus,95 biofilm-forming B. subtilis,96 B. circulans, B. laterosporus, B. lentus, and B. sphaericus97 in oral disease, mainly gingivitis and periodontitis. On the other hand, cryptic (having no definable origin) pyogenic infections of the central nervous system (CNS) secondary to dental affections have been reported in which B. subtilis and B. circulans are implicated.98 These cases indicate that, although dental and paradental infections seem to be a rare cause of intracerebral or intraspinal infections in otherwise healthy individuals, it is recommended that a dental focus should always be considered in the evaluation and treatment of these kinds of cryptic infections of the CNS. Meningoencephalitis and Meningitis. In addition to B. anthracis, to the best of our knowledge, only B. cereus has been reported to be implicated in cases of meningoencephalitis in preterm infants99 and neonates100 or in cases of meningitis in neonates101,102 and immunocompromised individuals.103,104 Soft-tissue Infection. To date, there have been two case reports that have implicated B. cereus in necrotizing fasciitis and myositis that mimick clostridial myonecrosis in immunocompromised patients.105,106 This is a severe infection of skin and soft tissue by B. cereus that has symptoms similar to clostridial gas gangrene: in both cases the infection is caused by a gram-positive rod, and gas production and muscles inflammation (myositis) could be observed. However, to select the appropriate therapy, it is very important to differentiate between the two kinds of infections. Indeed, the treatment of clostridial gas gangrene is usually based on penicillin, an antibiotic to which B. cereus is resistant because it produces β -lactamase. B. cereus has to be suspected as a pathogen when the skin and soft-tissue infection occurs with sepsis in immunocompromised patients; whereas, Clostridium have to

Bacillus

be considered when the infection occurs after trauma in otherwise healthy individuals. Another criterion for the differentiation between the two conditions is the site of infection: for clostridial gas gangrene, the primary site of inflammation is the muscle parenchyma, in contrast to B. cereus infection, where inflammation often develops from the subcutaneous tissues to muscles.106 Peritonitis. B. cereus,107,108 B. circulans,109 and B. liche­ niformis110 have also been implicated in peritonitis in patients  undergoing peritoneal dialysis. There is also one case report on B. cereus peritonitis after cesarean section.111

21.1.5  Diagnosis Techniques The diagnosis of a human illness is based on the combination of different clues including the symptoms of the illness, the history of the patient, the possible risk factors, and the detection and identification of the suspected agent and/or its pathogenicity factors. Conventional detection procedures of Bacillus human pathogens are often based in the first step on the isolation and identification of the bacterium on selective cultural media from clinical specimens based on biochemical, phenotypic, and physiological characteristics. Otherwise, detection of the whole microorganism or its pathogenecity factors could be performed by immunological assays. Nucleic acid–based approaches are, however, most-often applied due to their sensitivity and rapidity. 21.1.5.1  Conventional Diagnosis Techniques Fast and reliable identification of the pathogenic microorganism implicated in the illness is key to a timely and adequate treatment of a patient. Typically, B. anthracis can be identified from clinical specimens and differentiated from other Bacillus spp. by classical microbiological tests that are recommended by the World Health Organization (WHO) and by the Centers for Disease Control and Prevention (CDC). Like the other species of the B. cereus group, B. anthracis is positive to Gram staining and produces spores; however, it can be distinguished from the nonanthrax group (B. cereus and B. thuringiensis) by its capacity to produce a polypeptide capsule in vivo and in vitro when grown under suitable conditions, its sensitivity to gamma phage and penicillin G, and by the lack of motility and β-hemolytic activity on sheep or horse blood agar medium.57 In addition to blood agar medium, B. anthracis can be isolated on the selective polymyxin-lysozyme-ethylenediaminetetraacetic acid-thallium acetate (PLET) agar112 or on anthracis chromogenic agar (ACA).113 On PLET medium, B.  anthracis colonies appear small, white, domed, and circular. Anthracis chromogenic agar contains the chromogenic substrate 5-bromo-4-chloro-3-indoxyl-choline phosphate that yields teal colonies upon hydrolysis by phosphatidylcholine-specific phospholipase C (PC-PLC), also known as lecithinase C. Upon incubation at 35°C–37°C, B. cereus and B. thuringiensis colonies are dark teal-blue within 24 h; whereas, B. anthracis colonies need at least 48 h to develop

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this color because of the slower rate of synthesis of this enzyme as a result of the truncated plcR gene and PlcR regulator. However, the blood-agar medium is easier and more cost effective, and hence is recommended for the isolation of B. anthracis from samples not overloaded with background bacilli, such as clinical specimens. The selective media PLET and ACA are suitable for the isolation of B. anthracis by well trained and expert personal, from samples with high numbers of bacteria but taking into account that there are limitations.114 However, this classical identification procedure is timeconsuming and needs at least 24–48 h to be accomplished. In addition, there are several other challenges in the detection of B. anthracis: (i) not all the B. anthracis isolates fit all the above described characteristics, and atypical B.  anthracis isolates lacking pXO2 (and hence capsule) have been identified; (ii) pathogenic capsule-producing B. cereus strains harboring plasmids similar to those of B. anthracis have been identified in cases of human fatal pneumonia and from the carcasses of great apes in Africa115; and (iii) blood cultures do provide B. anthracis identification, whereas cultures from skin or lesions are not reliable as they yield positive results in only 60%–65% of cases, especially after antibiotic treatment.58 The limitations of culture-based diagnosis of pathogenic Bacillus spp. causing human illness make necessary the application of quicker and more specific approaches, especially for the detection of B. anthracis. Several antibodybased tests targeting spores, vegetative cells, and anthrax toxin proteins have been developed for the specific detection of B. anthracis.59,116–118 Recently, several other approaches targeting the B. anthracis capsular antigen detection by latex agglutination119 or the entire organism mediating phage bioluminescence120 have been reported. However, these antibody-based assays bear the limitation of sensitivity since low concentration of antigens could not be detected in the first steps of bacterial infection. 21.1.5.2  Molecular Diagnosis Techniques DNA isolation from clinical specimens is a critical step in molecular biology–based diagnosis techniques because of the presence of high levels of inhibitors and host DNA in relation to the amount of DNA from the pathogenic bacterium, especially in the first step of the infection. Also, safety is an important consideration for the personnel handling specimens that are suspected to contain infectious B. anthracis spores, which are known to be difficult to inactivate. Several extraction kits are commercially available for the rapid isolation of DNA to be used in nucleic acid–based detection assays. In particular, the UltraClean Microbial DNA Isolation Kit (Mo Bio Laboratories, Inc.) was shown to offer the best method for depleting viable B. anthracis spores from samples.121 Dauphin et al.122 reported that gamma irradiation of DNA preparations allowed inactivation of residual B. anthracis spores; their complete removal could be reached by centrifugal filtration with 0.1-micron filter unit.123

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21.2  METHODS 21.2.1  Sample Preparation Bacterial Isolation. Bacillus spp. are often isolated from clinical specimens on 5%–7% horse or sheep blood agar medium. After overnight incubation at 35°C, nonanthrax bacteria of the B. cereus group are gram-positive, rodshaped, spore formers, motile, and beta-hemolytic; whereas B. anthracis is nonmotile and nonhemolytic. B. anthracis is also sensitive to gamma phage and penicillin G. Media like PLET or ACA could also be used for the selective isolation of B. anthracis. 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 35°C. The cell pellet is washed once with 500 µL of sterile MilliQ water or TE buffer (pH = 8), centrifuged, resuspended in 100 µL of sterile MilliQ water or TE buffer, and boiled for 10–15 min. The boiled samples are centrifuged at 10,000 × 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 as well as from spores.121 The phenolchloroform extraction protocol is also useful to obtain pure high-molecular-weight genomic DNA.124 Total DNA from clinical specimens can be isolated using commercial DNA extraction kits, such as the ChargeSwitch gDNA Mini Bacteria Kit (Invitrogen); DNeasy Blood and Tissue kit (Qiagen); NucliSens Isolation Kit (Biomerieux); Puregene Genomic DNA Purification Kit (Qiagen); QIAamp DNA Blood Mini Kit (Qiagen); MagNA Pure LC (MPLC) microbiology kit MGrade (Roche Diagnostics GmbH); MagSi-DNA isolation kit for blood (Magna-Medics Diagnostics B.V.), and the UltraClean Microbial DNA Isolation Kit (Mo Bio Laboratories), following the manufacturers’ instructions. However, this step of sample processing may be hampered by the low concentration of the pathogen and the dominance of human DNA, especially in samples such as blood, cerebrospinal fluid, or in other primary sterile sites. Horz et al.125 developed two new kits, MolYsis® (Molzym GmbH & Co. KG, Bremen, Germany) and Pureprove® (SIRS-Lab GmbH, Jena, Germany) which is also known as Looxster® Universal kit, for the enrichment of bacterial DNA from blood samples that were also shown to be efficient using oral samples from plaque periodontitis and caries. Hansen et al.126 found that pretreatment of blood samples with MolYsis Basic prior to DNA isolation with MolYsis Complete Kit improves the detection of the DNA of pathogenic bacteria from blood samples allowing a detection limit of 50 CFU/mL in multiplex real-time PCR.

21.2.2  Detection Procedures Culture-dependent techniques for direct isolation and identification of Bacillus spp. based on traditional methods,

Molecular Detection of Human Bacterial Pathogens

looking to metabolism and physiology, is time consuming and laborious. Also, it is not possible to rely on bacterial culture for the diagnosis of cutaneous human anthrax, for which it has been reported that the bacterial culture– based diagnosis yields positive results only in 60%–65% of cases.58 Furthermore, the presence of atypical nonencapsulated B. anthracis strains and pathogenic capsule-producing B. cereus isolates represents a further challenge for a specific and accurate identification based on this phenotypic character. Thus, a combination of classical microbiological assays and nucleic acid–based methods is often necessary to confirm the identity of the infectious bacterial agent. Actually, there is a wide variety of molecular-based techniques for the detection of B. anthracis, including PCR, real-time PCR, and multiplex PCR. All these techniques are based on the specific amplification of the genes pag, lef, and cap. However, all these genetic determinants reside on plasmids; hence B. anthracis isolates that lack one of the plasmids can not be easily identified based on these techniques. For this reason, chromosomal-based B. anthracis–specific markers, such as BA813, rpoB, gyrA, gyrB, saspB, plcR, and BA5345 have been identified.51,127–129 Recently, Antwerpen et al.129 reported the development of a TaqMan real-time–based assay that targets the locus BA_5345, and that specifically detected B. anthracis, as was shown by testing 328 Bacillus strains. Within this sequence, they amplified a 96-bp large fragment by using the primers dhp61_183- 113F (5′-CGTAAGGACAATAAAAGCCGTTG T-3′), dhp61_183–208R (5′-CGATACAGACATTTATTGGG AACTACAC-3′) and TaqMan -probe dhp61_183–143T (5′-6FAM-TGCAATCGATGAGCTAATGAACAATGACCCT– TAMRA-3′). The TaqMan® PCR assay is performed in a 20  µL PCR-reaction mixture containing 1× ABITaqMan® mix (Applied Biosystems, Darmstadt, Germany); 0.9 µM primer dhp61_183–113F; 0.3 µM of primer dhp61_183–208R and 0.25 µM of labeled probe dhp61_183–143T; and 2 µL of template DNA. Amplification is carried out on the MX3000 Real-Time PCR System (Stratagene) with thermal conditions as follows: an initial denaturation step of 10 min at 95°C is followed by 45 cycles consisting of 15 s at 95°C and 1 min at 55°C. This assay is reported to have a detection limit of 12.7 copies/reaction. Another molecular assay that can be applied for the rapid and sensitive/quantitative detection of B. anthracis has been reported recently.130 This molecular assay is based on the combination of molecular beacons and real-time PCR technologies. Molecular beacons are highly specific nucleic acid probes that, when bound to their target DNA sequences, act as switches by emitting fluorescence as a result of the alteration of their conformation.130,131 The molecular beacon– based real-time PCR assay was performed by designing five molecular beacons targeting B. anthracis capA, capB, capC, lef, and pag alleles that were used in five uniplex assays for the detection of these genes in a broad range of samples.130 The assay is conducted in 25 μL reaction mixture containing 5.0 μL DNA, 4.5 μL distilled water, 12.5 μL Platinum Quantitative PCR SuperMix-UDG, 1.0 μL of one of the five

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Bacillus

molecular beacons and 1.0 μL (20 pmol/μL) of each of the forward and reverse primers. The assay is run on a 7900HT Real-Time PCR System (Applied Biosystems) with the following thermal cycling program: denaturation (94°C for 10 min), followed by 40 cycles of amplification (94°C denaturation for 15 s; 50°C annealing and data collection for 30 s; and 72°C polymerization for 30 s). During each cycle of amplification, the fluorescence emission in each sample is automatically recorded at 490 nm. Another innovative powerful gene-amplification technique that is becoming very popular and emerging as a simple and rapid diagnostic tool for the detection and identification of microbial diseases is the loop-mediated isothermal amplification (LAMP) that was described for the first time by Notomi et al.132 LAMP is based on the principle of strand-displacing DNA synthesis by the Bst DNA polymerase, with six distinct primers that recognize eight distinct sequences of a target gene. DNA amplification performed under isothermal conditions (60°C–65°C) using a heating block and/or water bath, is usually completed in less than 1 h. The gene-amplification products can be analyzed by agarose gel electrophoresis or by real-time monitoring of the turbidity of the reaction mixture in a turbidimeter. In this later case, it is also possible to quantify the gene copy number based on a standard curve generated from different concentrations of the gene copy number plotted against time of positivity. Moreover, gene amplification can be monitored by the naked eye either as turbidity or in the form of a color change when a fluorescent dsDNA intercalating dye is used. LAMP assays have been used for rapid detection of several pathogenic viruses, bacteria, and blood protozoa.133,134 Recently, a specific LAMP assay targeting the chromosomal sap gene that codifies for the S-layer constituent protein and cap and pag plasmid genes was designed by Kurosaki et  al.135 as a method for B. anthracis detection. Loopamp DNA Amplification Kit (Eiken Chemical, Tokyo, Japan) is used for performing LAMP reaction following the manufacturer’s instructions. Briefly, a 25 µL final volume reaction mixture containing 40 pmol of each forward inner primer (FIP) and a reverse inner primer (BIP); 5 pmol of each of the outer primers F3 and B3; 20 pmol of each of the two loop primers (LF and LB); 12.5 µL of 2× reaction mix; 1.0 µL of Bst DNA polymerase; and 5.0 µL of DNA sample are incubated at 63°C for 60 min. The LAMP products are analyzed on 1% agarose gel and ehidium bromide staining. Otherwise, for real-time monitoring of amplification, the reaction mixtures are incubated and observed by spectrophotometric analysis using a real-time turbidimeter (LA-200; Teramecs, Kyoto, Japan). The time of positivity observed through realtime LAMP assay is determined as the time at which the turbidity reached the threshold value fixed at 0.1, which is double the average turbidity value of negative controls in several replicates. The authors reported that this assay is 10- to 100-fold more sensitive than conventional PCR-based methods, allowing the detection of 10 fg of bacterial DNA per reaction and 3.6 CFU of bacterial spores in blood samples from experimentally infected mice.

21.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES Spores produced by Bacillus spp., other than B. anthracis, are ubiquitous in the environment owing to their resistance to harsh environmental conditions as well as to disinfection procedures. Their presence in food samples or clinical specimens has often been considered as a contamination. However, Bacillus spp. can cause serious and even lethal infections or intoxications not only in immunocompromised individuals or patients with other high-risk factors but also in otherwise healthy individuals. Conventional diagnostic procedures are based mainly on phenotypic and biochemical characteristics. For example, in the B. cereus group, the differentiation of the well-known opportunistic human pathogen B. cereus and the anthrax agent B. anthracis is based in part on the presence of a poly-gamma-D-glutamic acid capsule. However, both encapsulated B. cereus that has been implicated in fatal pneumonia and B. thuringiensis strains producing a capsule similar to the B. anthracis one136 have been discovered recently. This finding underlines the importance of using multiple tests for a reliable identification of isolates associated with human disease. Different Bacillus spp. outbreaks in hospital environments where the origin of the disseminated isolate was the reusable equipment, such as towels, gowns, and hand ventilator balloons, have been reported. Also, Bacillus spp. have been isolated from alcoholic cotton and detergents. These reported cases highlight the importance of using disposal or wrapping steamed towels and wrapping alcoholic cotton as preventive measures to avoid nosocomial bacteremia caused by these spore formers. They also underscore the importance of using professionally performed sterilization procedures for all reusable equipment and that should include eradication of spores. For this purpose, autoclaving is imperative and especially for reusable equipment in intensive care units and for material used by immunocompromised patients. The 70%–90% alcohol that is extensively used in hospitals is suitable as germicide but does not affect spores. On the other hand, regular cleaning of the filters of the air-conditioning system and the inlets and outlets of the air-ventilation system, as well as using the recommended concentration of sodium hypochlorite in the laundry process would be important measures to avoid nosocomial bacteremia by spore formers. Several molecular detection methods are actually available for B. anthracis and its pathogenecity factors, including the secreted anthrax toxins and the capsule, as well as 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. The conventional PCR bears the risk of false-positive results due to sample contamination. The multiplex PCR could be a good alternative for detecting different pathogens or different toxins of the same pathogenic bacterial species simultaneously. However, in some cases, it is possible to get false-negative results because of the presence of different primer pairs in the same reaction mix or different annealing

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temperatures. In addition, this method is qualitative and does 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 greenbased RT–PCRs could be applied. However, although highly sensitive, this technique could be biased by the presence of PCR inhibitors in clinical specimens. Also, the presence of dominant concentrations of human DNA in comparison to the DNA from the pathogenic bacterium in samples such as blood and cerebrospinal fluid, in particular, during the first step of the infection, can deeply affect the sensitivity of the PCR. Another practical drawback of RT–PCR is the necessity of expensive and sophisticated real-time platforms and reagents that are not available in most laboratories especially in developing countries. During this last decade, and since its description in 2000, LAMP technique has been largely used for the diagnosis of human pathogenic bacteria as well as viruses and protozoa. Only one recent report described the application of this assay for the diagnosis of B. anthracis in experimentally infected mice. The advantages of this technique consist of rapid amplification at one set temperature without the need of a thermal cycler, simple operation, and easy detection by a turbidimeter or naked eye. Also, combining LAMP assay with boiling extraction, it is possible to quickly detect very low copies of the target gene and hence to perform an efficient on-site diagnosis in less than 1 h without the need for culture growth or precise DNA extraction. Hence, LAMP can be considered an innovative technique that has potential applications for both clinical diagnosis and surveillance of infectious diseases in developing countries, without requiring sophisticated equipment or skilled personnel. It is, however, important to take into account that one of the major drawbacks of this assay is that designing the primers presents a difficulty because at least six distinct primers that recognize eight distinct sequences of a target gene are required for the performance of the assay. In conclusion, the most serious and important human pathogen within the genus Bacillus is B. anthracis. However, other species should also be considered in the diagnosis of septicemic infections, pneumonia, meningitis, endophthalmitis, and cutaneous infections. Among others, Bacillus cereus is being acknowledged as an emerging pathogen, especially in immunocompromised individuals. Also, although rare, capsule-producing pathogenic B. cereus strains with fatal anthrax-like pathology exist and should be considered, especially in patients with high-risk occupational factors, such as metal workers, and in particular, in zones where anthrax was endemic. As far as the detection of Bacillus spp. in clinical specimens, a combination of culture-based and molecular methods, including PCR amplification of the virulence factor encoding genes is necessary for the confirmation of the diagnosis and hence the choice of the appropriate therapy. To avoid misdiagnosis and unnecessary treatment, performing more than one set of cultures from clinical specimens is recommended before assuming that an isolate is a pathogen or dismissing a pathogen as a contaminant, which can

Molecular Detection of Human Bacterial Pathogens

cause complications and even mortality due to delay in the treatment.71

REFERENCES







1. Kim, K.M. et al., Characterization of Bacillus polyfermenticus KJS-2 as a Probiotic, J. Microbiol. Biotechnol., 19, 1013, 2009. 2. Sánchez, B. et al., Identification of surface proteins involved in the adhesion of a probiotic Bacillus cereus strain to mucin, Microbiology, 155, 1708, 2009. 3. Schwartz, M., Dr. Jekyll, and Mr. Hyde, A short history of anthrax, Mol. Aspects. Med., 30, 347, 2009. ‌ 4. Shimoyama, R. et al., Contamination of Bacillus cereus in an apheresis-derived platelet concentrate, J. Clin. Apheresis, 12, 103, 1997. 5. Pinna, A. et al., Bacillus cereus keratitis associated with contact lens wear, Ophthalmology, 108, 1830, 2001. 6. Störmer, M. et al., Spore-forming organisms in platelet concentrates: A challenge in transfusion bacterial safety, Transfus. Med., 18, 371, 2008. 7. Ohsaki, Y. et al., Undetected Bacillus pseudo-outbreak after renovation work in a teaching hospital, J. Infect., 54, 617, 2007. 8. Dohmae, S. et al., Bacillus cereus nosocomial infection from reused towels in Japan, J. Hosp. Infect., 69, 361, 2008. 9. Kalpoe, J.S. et al., Dissemination of Bacillus cereus in a paediatric intensive care unit traced to insufficient disinfection of reusable ventilator air-flow sensors, J. Hosp. Infect., 68, 341, 2008. 10. Kuroki, R. et al., Nosocomial bacteremia caused by biofilmforming Bacillus cereus and Bacillus thuringiensis, Intern. Med., 48, 791, 2009. 11. Daffonchio, D., Cherif, A., and Borin, S., Homoduplex and heteroduplex polymorphisms of the amplified ribosomal 16S23S internal transcribed spacers describe genetic relationships in the “Bacillus cereus group,” Appl. Environ. Microbiol., 66, 5460, 2000. 12. Cherif, A. et al., Genetic relationship in the “Bacillus cereus group” by rep-PCR fingerprinting and sequencing of a Bacillus anthracis-specific rep-PCR fragment, J. Appl. Microbiol., 94, 1108, 2003. 13. Dierick, K. et al., Fatal family outbreak of Bacillus cereusassociated food poisoning, J. Clin. Microbiol., 43, 4277, 2005. 14. Raddadi, N. et al., Bacillus. In Liu, D. (Editor), Molecular Detection of Food-borne Pathogens, 1st ed., chapter 10, Taylor & Francis Group, New York, 2009. 15. Baron, F. et al., Isolation and characterization of a psychrotolerant toxin producer, Bacillus weihenstephanensis, in liquid egg products, J. Food Prot., 70, 2782, 2007. 16. Nakamura, L.K., DNA relatedness among Bacillus thuringiensis serovars, Int. J. Syst. Bacteriol., 44, 125, 1994. 17. Nakamura, L.K., and Jackson, M.A., Clarification of the taxonomy of Bacillus mycoides, Int. J. Syst. Bacteriol., 45, 46, 1995. 18. Nakamura, L.K., Bacillus pseudomycoides sp. nov., Int. J. Syst. Bacteriol., 48, 1031, 1998. 19. Ash, C. et al., Phylogenetic heterogeneity of the genus Bacillus revealed by comparative analysis of small-subunit-ribosomal RNA sequences, Lett. Appl. Microbiol., 13, 202, 1991. 20. Ash, C. et al., Comparative analysis of Bacillus anthracis, Bacillus cereus, and related species on the basis of reverse transcriptase sequencing of 16S rRNA, Int. J. Syst. Bacteriol., 41, 343, 1991.

Bacillus 21. Ash, C., and Collins, M.D., Comparative analysis of 23S ribosomal RNA gene sequences of Bacillus anthracis and emetic Bacillus cereus determined by PCR-direct sequencing, FEMS Microbiol. Lett., 94, 75, 1992. 22. Brousseau, R. et al., Arbitrary primer polymerase chain reaction, a powerful method to identify Bacillus thuringiensis serovars and strains, Appl. Environ. Microbiol., 59, 114, 1993. 23. Carlson, C.R., Grønstad, A., and Kolstø, A.-B., Physical maps of the genomes of three Bacillus cereus strains, J. Bacteriol., 174, 3750, 1992. 24. Carlson, C.R., and Kolstø, A.-B., A complete physical map of a Bacillus thuringiensis chromosome, J. Bacteriol., 175, 1053, 1993. 25. Carlson, C.R., Caugant, D.A., and Kolstø, A.-B., Genotypic diversity among Bacillus cereus and Bacillus thuringiensis strains, Appl. Environ. Microbiol., 60, 1719, 1994. 26. Carlson, C.R., Johansen, T., and Kolstø, A.-B., The chromosome map of Bacillus thuringiensis subsp. canadensis HD224 is highly similar to that of the Bacillus cereus type strain ATCC 14579, FEMS Microbiol. Lett., 141, 163, 1996. 27. Daffonchio, D. et al., Restriction Site Insertion-PCR (RSIPCR) for rapid discrimination and typing of closely related microbial strains, FEMS Microbiol. Lett., 180, 77, 1999. 28. Daffonchio, D. et al., A randomly amplified polymorphic DNA marker specific for the Bacillus cereus group is diagnostic for Bacillus anthracis, Appl. Environ. Microbiol., 65, 1298, 1999. 29. Kim, Y.R., Czajka, J., and Batt, C.A., Development of a fluorogenic probe-based PCR assay for detection of Bacillus cereus in nonfat dry milk, Appl. Environ. Microbiol., 66, 1453, 2000. 30. Schraft, H., and Griffith, M.W., Specific oligonucleotide primers for detection of lecitinase-positive Bacillus spp. by PCR, Appl. Environ. Microbiol., 61, 98, 1995. 31. Schraft, H. et al., Epidemiological typing of Bacillus spp. isolated from food, Appl. Environ. Microbiol., 62, 4229, 1996. 32. Margulis, L. et al., The Arthromitus stage of Bacillus cereus: Intestinal symbionts of animals, Proc. Natl. Acad. Sci. USA, 95, 1236, 1998. 33. Jensen, G.B. et al., The hidden lifestyles of Bacillus cereus and relatives, Environ. Microbiol., 5, 631, 2003. 34. Swiecicka, I., and Mahillon J., Diversityof commensal Bacillus cereus sensu latoisolated from the commonsowbug (Porcellio scaber, Isopoda), FEMS Microbiol. Ecol., 56, 132, 2006. 35. Ciammaruconi, A. et al., Fieldable genotyping of Bacillus anthracis and Yersinia pestis based on 25-loci multi locus VNTR analysis, BMC Microbiol., 8, 21, 2008. 36. Ivanova, N. et al., Genome sequence of Bacillus cereus and comparative analysis with Bacillus anthracis, Nature, 423, 87, 2003. 37. Gohar, M. et al., A comparative study of Bacillus cereus, Bacillus thuringiensis and Bacillus anthracis extracellular proteomes, Proteomics, 5, 3696, 2005. 38. Keim, P. et al., Molecular evolution and diversity in Bacillus anthracis as detected by amplified fragment length polymorphism markers, J. Bacteriol., 179, 818, 1997. 39. Radnedge, L. et al., Genome differences that distinguish Bacillus anthracis from Bacillus cereus and Bacillus thuringiensis, Appl. Environ. Microbiol., 69, 2755, 2003. 40. Read, T.D. et al., The genome sequence of Bacillus anthracis Ames and comparison to closely related bacteria, Nature, 423, 81, 2003.

227 41. Sanz, P. et al., Detection of Bacillus anthracis spore germination in vivo by bioluminescence imaging, Infect. Immun., 76, 1036, 2008. 42. Helgason, E. et al., Multilocus sequence typing scheme for bacteria of the Bacillus cereus group, Appl. Environ. Microbiol., 70, 191, 2004. 43. Zwick, M.E. et al., Microarray-based resequencing of multiple Bacillus anthracis isolates, Genome Biol., 6, R10, 2005. 44. Keim, P. et al., Multiple-locus variable-number tandem repeat analysis reveals genetic relationships within Bacillus anthracis, J. Bacteriol., 182, 2928, 2000. 45. Read, T.D. et al., Comparative genome sequencing for discovery of novel polymorphisms in Bacillus anthracis, Science, 296, 2028, 2002. 46. Gierczyn´ski, R. et al., Intriguing diversity of Bacillus anthracis in eastern Poland—the molecular echoes of the past outbreaks, FEMS Microbiol. Lett., 239, 235, 2004. 47. Merabishvili, M. et al., Diversity of Bacillus anthracis strains in Georgia and of vaccine strains from the former Soviet Union, Appl. Environ. Microbiol., 72, 5631, 2006. 48. Prüss, B.M. et al., Correlation of 16S ribosomal DNA signature sequences with temperature-dependent growth rates of mesophilic and psychrotolerant strains of the Bacillus cereus group, J. Bacteriol., 181, 2624, 1999. 49. Van Ert, M.N. et al., Global genetic population structure of Bacillus anthracis, PLoS ONE, 2, e461, 2007. 50. Prüss, B.M. et al., Correlation of 16S ribosomal DNA signature sequences with temperature-dependent growth rates of mesophilic and psychrotolerant strains of the Bacillus cereus group, J. Bacteriol., 181, 2624, 1999. 51. 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. 52. Nicholson, W.L. et al., Resistance of Bacillus endospores to extreme terrestrial and extraterrestrial environments, Microbiol. Mol. Biol. Rev., 64, 548, 2000. 53. Cybulski, R.J. et al., Four superoxide dismutases contribute to Bacillus anthracis virulence and provide spores with redundant protection from oxidative stress, Infect. Immun., 77, 274, 2009. 54. Hugh-Jones, M., and Blackburn, J. The ecology of Bacillus anthracis, Mol. Aspects. Med., 30, 356, 2009. ‌ 55. Karahocagil, M.K. et al., Cutaneous anthrax in Eastern Turkey: A review of 85 cases, Clin. Exp. Dermatol., 33, 406, 2008. 56. Ozdemir, H. et al., Anthrax of the gastrointestinal tract and oropharynx: CT findings, Emerg. Radiol., DOI 10.1007/ s10140–009-0821-y, 2009. ‌ 57. World Health Organization, Anthrax in Humans and Animals, 4th ed., WHO, Geneva. Available from http://www.who.int/ csr/resources/publications/ anthrax_webs.pdf, 2008. 58. Dixon, T. et al., Anthrax, N. Engl. J. Med., 341, 815, 1999. 59. Edwards, K. et al., Bacillus anthracis: Toxicology, epidemiology and current rapid-detection methods, Anal. Bioanal. Chem., 384, 73, 2006. 60. Drysdale, M., Bourgogne, A., and Koehler, T.M. Transcriptional analysis of the Bacillus anthracis capsule regulators, J. Bacteriol., 187, 5108, 2005. 61. Fouet, A., and Mock, M., Regulatory networks for virulence and persistence of Bacillus anthracis, Curr. Opin. Microbiol., 9, 160, 2006. 62. Fang, F.C., Antimicrobial reactive oxygen and nitrogen species: Concepts and controversies, Nat. Rev. Microbiol., 2, 820, 2004.

228 63. van Sorge, N.M. et al., Anthrax toxins inhibit neutrophil signaling pathways in brain endothelium and contribute to the  pathogenesis of meningitis, PLoS One, 3, doi:10.1371/ journal.pone.0002964, 2008. 64. Sejvar, J.J., Tenover, F.C., and Stephens, D.S. Management of anthrax meningitis, Lancet. Infect. Dis., 5, 287, 2005. 65. Simonson, T.S. et al., Bacillus anthracis in China and its relationship to worldwide lineages, BMC Microbiol., 9, 71, 2009. 66. Doganay, M., and Metan, G., Human Anthrax in Turkey from 1990 to 2007, Vector-born. Zoonot., 9, DOI: 10.1089/ vbz.2008.0032, 2009. 67. Lanska, D., Anthrax meningoencephalitis, Neurology, 59, 327, 2002. 68. Narayan, S.K. et al., Anthrax meningoencephalitis—declining trends in an uncommon but catastrophic CNS infection in rural Tamil Nadu, South India, J. Neurol. Sci., 281, 41, 2009. 69. Weber, D.J. et al., Clinical significance of Bacillus species isolated from blood cultures, South. Med. J., 82, 705, 1989. 70. Lee, C.C. et al., Clinical significance of potential contaminants in blood cultures among patients in a medical center, J. Microbiol. Immunol. Infect., 40, 438, 2007. 71. Ashkenazi-Hoffnung, L. et al., Seasonality of Bacillus species isolated from blood cultures and its potential implications, Am. J. Infect. Control, 37, 495, 2009. 72. Van Der Zwet, W.C. et al., Outbreak of Bacillus cereus infections in a neonatal intensive care unit traced to balloons used in manual ventilation, J. Clin. Microbiol., 38, 4131, 2000. 73. Hernaiz, C. et al., Nosocomial bacteremia and catheter infection by Bacillus cereus in an immunocompetent patient, Clin. Microbiol. Infect., 9, 973, 2003. 74. Ghelardi, E. et al., Bacillus thuringiensis pulmonary infection: Critical role for bacterial membrane-damaging toxins and host neutrophils, Microbes Infect., 9, 591, 2007. 75. Ozkocaman, V. et al., Bacillus spp. among hospitalized patients with haematological malignancies: Clinical features, epidemics and outcomes, J. Hosp. Infect., 64, 169, 2006. 76. Lépine, A. et al., Bacillus licheniformis septicemia in a verylow-birth-weight neonate: A case report, Infection, 37, 156, 2009. 77. Blue, S.R., Singh, V.R., and Sanbolle, M.A., Bacillus licheniformis bacteraemia: Five cases associated with indwelling central venous catheters, Clin. Infect. Dis., 20, 629, 1995. 78. Haymore, B.R. et al., A case of persistent Bacillus pumilis bacteremia associated with cholangitis, J. Infect., 52, 154, 2006. 79. Bentur, H.N., Dalzell, A.M., and Riordan, F.A.I. Central venous catheter infection with Bacillus pumilus in an immunocompetent child: A case report, Ann. Clin. Microbiol. Antimicrob., 6, 12, 2007. 80. Ramakrishnan, R. et al., Microbiological profile of culture proven cases of exogenous and endogenous endophthalmitis: A 10-year retrospective study, Eye, 23, 945, 2009. 81. Ramos-Esteban, J.C. et al., Bacillus megaterium delayed onset lamellar keratitis after LASIK, J. Refract. Surg., 22, 309, 2006. 82. Callegan, M.C. et al., Virulence factor profiles and antimicrobial susceptibilities of ocular bacillus isolates, Curr. Eye. Res., 31, 693, 2006. 83. Kopel, A.C., Carvounis, P.E., and Holz E.R. Bacillus cereus endophthalmitis following intravitreous bevacizumab injection, Ophthalmic. Surg. Lasers. Imaging., 39, 153, 2008. 84. Callegan, M.C. et al., Bacillus endophthalmitis: Roles of bacterial toxins and motility during infection, Invest. Ophthalmol. Vis. Sci., 46, 3233, 2005.

Molecular Detection of Human Bacterial Pathogens 85. Ramadan, R.T. et al., A role for tumor necrosis factor-alpha (TNFα) in experimental Bacillus cereus endophthalmitis pathogenesis, Invest. Ophthalmol. Vis. Sci., 49, 4482, 2008. 86. Moyer, A.L. et al., Bacillus cereus-induced permeability of the blood-ocular barrier during experimental endophthalmitis, Invest. Ophthalmol. Vis. Sci., 50, 3783, 2009. 87. Mahler, H. et al., Fulminant liver failure in association with the emetic toxin of Bacillus cereus, N. Engl. J. Med., 336, 1142, 1997. 88. Latsios, G. et al., Liver abscess due to Bacillus cereus: A case report, Clin. Microbiol. Infect., 9, 1234, 2003. 89. Hoffmaster, A.R. et al., Identification of anthrax toxin genes in a Bacillus cereus associated with an illness resembling inhalation anthrax, Proc. Natl. Acad. Sci. USA, 101, 8449, 2004. 90. Miller, J.M. et al., Fulminating bacteremia and pneumonia due to B. cereus, J. Clin. Microbiol., 35, 504, 1997. 91. Avashia, S.B. et al., Fatal pneumonia among metalworkers due to inhalation exposure to Bacillus cereus containing Bacillus anthracis toxin genes, Clin. Infect. Dis., 44, 414, 2007. 92. Hoffmaster, A.R. et al., Characterization of Bacillus cereus isolates associated with fatal pneumonias: strains are closely related to Bacillus anthracis and harbor B. anthracis virulence genes, J. Clin. Microbiol., 44, 3352, 2006. 93. Katsuya, H. et al., A patient with acute myeloid leukemia who developed fatal pneumonia caused by carbapenem-resistant Bacillus cereus, J. Infect. Chemother., 15, 39, 2009. 94. Helgason, E. et al., Genetic structure of population of Bacillus cereus and B. thuringiensis isolates associated with periodontitis and other human infections, J. Clin. Microbiol., 38, 1615, 2000. 95. Johnson, B.T. et al., Extracellular proteolytic activities expressed by Bacillus pumilus isolated from endodontic and periodontal lesions, J. Med. Microbiol., 57, 643, 2008. 96. Yamane, K. et al., Identification and characterization of clinically isolated biofilm-forming gram-positive rods from teeth associated with persistent apical periodontitis, J. Endod., 35, 347, 2009. 97. Sunde, P.T. et al., Microbiota of periapical lesions refractory to endodontic therapy, J. Endod., 28, 304, 2002. 98. Ewald C., Kuhn, S., and Kalff, R. Pyogenic infections of the central nervous system secondary to dental affection—a report of six cases, Neurosurg. Rev., 29, 163, 2006. 99. Lequin, M.H. et al., Bacillus cereus meningoencephalitis in preterm infants: Neuroimaging characteristics, Am. J. Neuroradiol., 26, 2137, 2005. 100. Manickam, N., Knorr, A., and Muldrew, K.L. Neonatal meningoencephalitis caused by Bacillus cereus, Pediatr. Infect. Dis. J., 27, 843, 2008. 101. Evreux, F. et al., A case of fatal neonatal Bacillus cereus meningitis, Arch. Pediatr., 14, 365, 2007. 102. Lebessi, E. et al., Bacillus cereus meningitis in a term neonate, J. Matern. Fetal. Neonatal. Med., 22, 458, 2009. 103. de Almeida, S.M. et al., Fatal Bacillus cereus meningitis without inflammatory reaction in cerebral spinal fluid after bone marrow transplantation, Transplantation, 76, 1533, 2003. 104. Haase, R. et al., Successful treatment of Bacillus cereus meningitis following allogenic stem Cell transplantation, Pediatr. Transplant., 9, 338, 2005. 105. Meredith, F.T. et al., Bacillus cereus necrotizing cellulitis mimicking clostridial myonecrosis: Case report and review of the literature, Scand. J. Infect. Dis., 29, 528, 1997. 106. Sada, A., Necrotizing fasciitis and myonecrosis “synergistic necrotizing cellulitis” caused by Bacillus cereus, J. Dermatol., 36, 423, 2009.

Bacillus 107. Pinedo, S., Bos, A.J., and Siegert, C.E. Relapsing Bacillus cereus peritonitis in two patients on peritoneal dialysis, Perit. Dial. Int., 22, 424, 2002. 108. Monteverde, M.L. et al., Relapsing Bacillus cereus peritonitis in a pediatric patient on chronic peritoneal dialysis, Perit. Dial. Int., 26, 715, 2006. 109. Berry, N. et al., Bacillus circulans peritonitis in a patient treated with CAPD, Perit. Dial. Int., 24, 488, 2004. 110. Park, D.J. et al., Relapsing Bacillus licheniformis peritonitis in a continuous ambulatory peritoneal dialysis patient, Nephrology, 11, 21, 2006. 111. Shimoni, Z. et al., Bacillus cereus peritonitis after Cesarean section, Surg. Infect., 9, 105, 2008. 112. Knisely, R.F., Selective medium for Bacillus anthracis, J. Bacteriol., 92, 784, 1966. 113. Juergensmeyer, M.A. et al., A selective chromogenic agar that distinguishes Bacillus anthracis from Bacillus cereus and Bacillus thuringiensis, J. Food. Protect., 69, 2002, 2006. 114. Marston, C.K. et al., Evaluation of two selective media for the isolation of Bacillus anthracis, Lett. Appl. Microbiol., 47, 25, 2008.‌ 115. Klee, S.R. et al., Characterization of Bacillus anthracis-like bacteria isolated from wild great apes from Cote d’Ivoire and Cameroon, J. Bacteriol., 188, 5333, 2006. 116. Hao, R. et al., Rapid detection of Bacillus anthracis using monoclonal antibody functionalized QCM sensor, Biosens. Bioelectron. 24, 1330, 2009. 117. Tang, S. et al., Detection of anthrax toxin by an ultrasensitive immunoassay using europium nanoparticles, Clin. Vaccine. Immunol., 16, 408, 2009. 118. Duriez, E. et al., Femtomolar detection of the anthrax edema factor in human and animal plasma, Anal. Chem., 81, 5935, 2009. 119. AuCoin, D.P. et al., Rapid detection of the poly-γ-D-glutamic acid capsular antigen of Bacillus anthracis by latex agglutination, Diagn. Microbiol. Infect. Dis., 64, 229, 2009. 120. Schofield, D.A., and Westwater, C., Phage-mediated bioluminescent detection of Bacillus anthracis, J. Appl. Microbiol., 107, 1468, 2009. ‌ 121. Dauphin, L.A., Moser, B.D., and Bowen, M.D. Evaluation of five commercial nucleic acid extraction kits for their ability to inactivate Bacillus anthracis spores and comparison of DNA yields from spores and spiked environmental samples, J. Microbiol. Methods, 76, 30, 2009. 122. Dauphin, L.A. et al., Gamma irradiation can be used to inactivate Bacillus anthracis spores without compromising the

229 sensitivity of diagnostic assays, Appl. Environ. Microbiol., 74, 4427, 2008. 123. Dauphin, L.A., and Bowen, M.D., A simple method for the rapid removal of Bacillus anthracis spores from DNA preparations, J. Microbiol. Methods, 76, 212, 2009. 124. Sambrook, J., Fritsch, E.F., and Maniatis, T., Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, 1989. 125. Horz, H.P. et al., Selective isolation of bacterial DNA from human clinical specimens, J. Microbiol. Methods, 72, 98, 2008. 126. Hansen, W.L., Bruggeman, C.A., and Wolffs, P.F. Evaluation of new preanalysis sample treatment tools and DNA isolation protocols to improve bacterial pathogen detection in whole blood, J. Clin. Microbiol., 47, 2629, 2009. 127. Ramisse, V. et al., The Ba813 chromosomal DNA sequence effectively traces the whole Bacillus anthracis community, J. Appl. Microbiol., 87, 224, 1999. 128. Qi, Y. et al., Utilization of the rpoB gene as a specific chromosomal marker for real-time PCR detection of Bacillus anthracis, Appl. Environ. Microbiol., 67, 3720, 2001. 129. Antwerpen, M.H. et al., Real-time PCR system targeting a chromosomal marker specific for Bacillus anthracis, Mol. Cell. Probes, 22, 313, 2008. 130. Hadjinicolaou, A.V., and Victoria, L., Use of molecular beacons and multi-allelic real-time PCR for detection of and discrimination between virulent Bacillus anthracis and other Bacillus isolates, J. Microbiol. Methods, 78, 45, 2009. 131. Li, Y. et al., Molecular beacons: An optimal multifunctional biological probe, Biochem. Biophys. Res. Comm., 373, 457, 2008. 132. Notomi, T. et al., Loop-mediated isothermal amplification of DNA, Nucleic Acids Res., 28, E63, 2000. 133. Mori, Y. et al., Detection of loop-mediated isothermal amplification reaction by turbidity derived from magnesium pyrophosphate formation, Biochem. Biophys. Res. Comm., 289, 150, 2001. 134. Parida, M. et al., Loop mediated isothermal amplification (LAMP): A new generation of innovative gene amplification technique; perspectives in clinical diagnosis of infectious diseases, Rev. Med. Virol., 18, 407, 2008. 135. Kurosaki, Y. et al., A simple and sensitive method for detection of Bacillus anthracis by loop-mediated isothermal amplification, J. Appl. Microbiol., 107, 1947, 2009. ‌ 136. Cachat, E. et al., A Bacillus thuringiensis strain producing a polyglutamate capsule resembling that of Bacillus anthracis, FEMS Microbiol. Lett., 285, 220, 2008.

22 Enterococcus Teresa Semedo-Lemsaddek, Paula Lopes Alves, Rogério Tenreiro, and Maria Teresa Barreto Crespo CONTENTS 22.1 Introduction...................................................................................................................................................................... 231 22.1.1 Classification and Morphology............................................................................................................................. 231 22.1.2 Clinical Features, Epidemiology, and Pathogenesis............................................................................................. 231 22.1.3 Diagnosis.............................................................................................................................................................. 233 22.1.3.1 Phenotypic Techniques.......................................................................................................................... 233 22.1.3.2 Molecular Techniques............................................................................................................................ 233 22.2 Methods............................................................................................................................................................................ 241 22.2.1 Sample Preparation............................................................................................................................................... 241 22.2.2 Detection Procedures............................................................................................................................................ 241 22.2.2.1 Semiautomatic and Automatic Identification......................................................................................... 241 22.2.2.2 Multiplex PCR for the Detection of Enterococcus Species................................................................... 242 22.3 Conclusions and Future Perspectives............................................................................................................................... 243 Acknowledgments...................................................................................................................................................................... 243 References.................................................................................................................................................................................. 243

22.1  INTRODUCTION 22.1.1  Classification and Morphology Enterococci are mainly viewed as commensal bacteria inhabiting the gastrointestinal tract of humans and animals, but in fact, these microorganisms have been recognized as potentially pathogenic for humans for more than a century. In Thiercelin’s1 original description, these bacteria had been isolated from patients with enteritis, appendicitis, and meningitis.2 The classification and nomenclature of the genus Enterococcus has raised interest and constant discussion over the years, ever since Thiercelin first proposed the name “enterocoques” in 1902. In 1970, Kalina3 proposed that Streptococcus faecalis and S. faecium should be renamed as Enterococcus faecalis and E. faecium, respectively. These proposals were not officially recognized for many years, and the genus Enterococcus did not appear in the Approved List of Bacterial Names until the 1980s. In the 1980s, Schleifer and Kilpper-Bälz4 resurrected the genus Enterococcus after performing DNA–DNA and DNA–rRNA hybridizations. Other studies carried out in the same decade, including 16S rRNA sequencing,5 also demonstrated that enterococci form a separate genus. Analysis of these molecular data divided the streptococci sensu lato into three genera: (i) Streptococcus 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, Kohler6 performed an extensive review of the publications that lead to each of the species currently accepted and included in TOBA (Taxonomic Outline of the Bacteria and Archaea),7 based on a comprehensive taxonomy using 16S rRNA gene phylogeny. Members of the genus Enterococcus are gram-positive ovoid cells that appear singly, in pairs, or in short chains; are 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 10ºC 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%.8,9 However, not all enterococci exhibit these characteristics; several authors have analyzed and reported differences in physiological properties among different species and strains belonging to the same species,10–15 making a reliable identification at species level difficult.

22.1.2  Clinical Features, Epidemiology, and Pathogenesis Although enterococci have traditionally been regarded as low-grade pathogens, in the last decades they have emerged 231

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as an increasingly important cause of nosocomial infections, mainly in patients with underlying disease. All kinds of severe immunosupression (e.g., oncology, hematology, nephrology, or transplantation units), long hospital stays,16 or the presence of urinary or vascular catheters constitute notable risk factors.17 The infections most frequently caused by enterococci are urinary tract infections (UTIs),18 followed by intraabdominal/ intrapelvic abscesses or postsurgery wound and bloodstream infections (BSIs).19 Enterococci are also infrequent causes of central nervous system (CNS) infections, neonatal infections, respiratory tract infections, osteomyelitis, or cellulitis.20 The importance of enterococci in the hospital setting is increasing every year. Enterococci are the third most common cause of nosocomial blood infections in the United States,21 and in Europe, the fourth (http://www.earss.rivm. nl/). Regarding urinary tract infections, they rank second in both the United States and Europe.22,23 Although more than thirty species have been identified,7 only two are responsible for the majority of human infections. The species E. faecalis and E. faecium account for 80% and 20% of the enterococcal infections, respectively. The proportion of E. faecium infections is increasing, and other species are being more frequently isolated from clinical samples, namely, E. gallinarum, E. casseliflavus, and E. raffinosus.22,24,25 The inherent tolerance of enterococci to unfavorable conditions allows them to survive for long periods in hospital settings, even on dry surfaces, and medical staff and instruments can be responsible for the transmission between hospitalized patients.17,26–28 These characteristics contribute to an increased risk of enterococcal nosocomial infections and emphasize the importance of bacterial control in the health care environment. Enterococcal infections are an important health concern, and as nosocomial pathogens, they have a direct and significant economic impact by prolonging hospital stays and requiring additional therapeutic treatments.29 Another important factor contributing to the increasing number of enterococcal infections is the uncontrolled use of antimicrobial agents, which enhances the selective pressure for bacteria with natural and/or acquired antibiotic resistances, such as enterococci. Administration of antimicrobial agents that have little effect on enterococci results in the elimination of many endogenous microbiota and may open new niches for the overgrowth of pathogenic strains. Enterococci have intrinsic resistance to several antimicrobial agents (e.g., cephalosporins, lincosamides, many β-lactams and low levels of aminoglycosides); frequently acquire plasmids and transposons carrying antimicrobial resistance genes (e.g., chloramphenicol, erythromycin, tetracyclin, and glycopeptides such as vancomycin) that can subsequently be transferred to other strains sharing the same ecological niche.2,16,26,28 Vancomycin resistance gains increased importance because this antimicrobial is considered a last resource treatment for many human infections. Vancomycinresistant enterococci (VRE) were first detected in Europe

Molecular Detection of Human Bacterial Pathogens

(United  Kingdom, and France) in 1988, and in the following year in the United States.30 In the last decades, the incidence of VRE rose, mainly due to E. faecium isolates, and vancomycin-resistant strains have already been reported in countries throughout the world, such as Australia, Belgium, Canada, Denmark, Germany, Italy, Malaysia, Netherlands, Spain, Sweden, and Portugal.31 While the increase of VRE in the United States is attributed to the uncontrolled usage of antimicrobials in hospitals,31 in Europe the reservoir of resistant strains seems to be linked with the use of avoparcin as a feed supplement for various livestock, and contamination via the food chain has been suggested.32–35 The usage of pulsed field gel electrophoresis has also demonstrated that acute outbreaks are typically related to a single VRE clone or a limited number of clones.36–38 Asymptomatic patients with VRE colonizing the gastrointestinal tract are usually the reservoir, and the dissemination between patients is mainly attributed to the indirect contact of the hands of health care workers. Surprisingly, little is known about the factors that contribute to the ability of enterococci to cause infection. In E. faecalis, several putative virulence factors have been described over the last years, involved either in the attachment to host cells or to extracellular matrix proteins, or implicated in cell and tissue damage. For gastrointestinal commensals such as enterococci, adhesins that promote binding to host cell receptors are expected to play an important role in the establishment and maintenance of colonization.39 The enterococcal surface protein (Esp) is a high molecular weight extracellular surface protein of unknown function whose frequency is increased among infection-derived isolates.40 This protein contributes to colonization and persistence of enterococci in the urinary tract41 and also to biofilm formation on abiotic surfaces.42 Aggregation substance (AS) is a pheromone-inducible surface protein that mediates the binding of donor cells to plasmid free recipients.43,44 This protein is essential for high-efficiency conjugation of sex pheromone plasmids, contributing to the dissemination of antimicrobial resistance and virulence determinants. Different functions have been attributed to AS in addition to bacterial cell aggregation, including adherence to host tissues45,46 and internalization,47 adhesion to fibrin and increased cell-surface hydrophobicity,48 resistance to killing by polymorphonuclear leukocytes and macrophages,46,49 and increased vegetation size in experimental endocarditis50 and endophthalmitis models.51 Another adhesion-associated protein was identified while analyzing serum from patients with E. faecalis endocarditis (named EfaA for E. faecalis antigen A).52,53 Regarding cell and tissue damage, the first virulence factor to be studied was cytolysin, a posttranslationally modified protein toxin that causes a β-hemolytic reaction on certain blood erythrocytes and also possesses bactericidal activity against a broad range of gram-positive bacteria.54–56 Gelatinase is an extracellular zinc-endopeptidase capable of hydrolysing gelatin, collagen, casein, and other small biologically active peptides, and its potential contribution to virulence has been suggested in several studies.57–60

Enterococcus

As regards E. faecium, only three putative virulence genes have been identified; they encode an enterococcal surface protein (espefm), a putative hyaluronidase (hyl)61,62 and an endocarditis-related adhesin EfaAfm.52,53 The putative virulence factors described for the genus Enterococcus can either be encoded by mobile genetic elements, such as plasmids or transposons, or by the bacterial chromosome. These genomic region clustering genes that contribute, directly or indirectly, to the pathogenic potency of the harboring microorganism are termed pathogenicity islands or PAIs.63 The generalized sequencing of genomes revealed that PAIs are much more widespread than previously thought.63 Regarding the genus Enterococcus, two PAIs have been described: one in a vancomycin-resistant strain of E. faecalis that caused a nosocomial outbreak,64 and the other in a group of nosocomial strains of E. faecium.65 More recently, the worldwide dissemination of the E. faecalis PAI among isolates from diverse locations and origins was demonstrated.66

22.1.3  Diagnosis 22.1.3.1  Phenotypic Techniques Detection and reliable identification, especially regarding VRE, is of major concern since VRE leaves very few options for the treatment of enterococcal infections.16,28 At the present time in Europe, VRE infections are increasing; meanwhile, in the United States, colonization of hospitalized patients is endemic in many tertiary care hospitals and increasingly causes infections.16 Conventional culture methods remain the most reliable and accurate techniques for the detection of bacterial pathogens such as enterococci. 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 neither a phylogenetically nor phenotypically coherent and homogeneous genus, manual or automated systems based on phenotypic characteristics do not always allow the correct strain identification; therefore these problems assume great importance.9 Another disadvantage of the standard culture methods is their lack of ability to detect a substantial proportion of the bacterial population that is nonculturable. These viable but nonculturable (VBNC) microorganisms demonstrate metabolic activity,67,68 maintain their pathogenicity69,70 and their antibiotic resistance traits,71 can express72 and exchange genes,73 and are able to reestablish division upon restoration of favorable conditions.74,75 It has been previously demonstrated that E. faecalis can enter the VBNC state,72 thus representing a risk in hospital settings and emphasizing the importance of molecular

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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. 22.1.3.2  Molecular Techniques Nucleic acid–based identification and detection methods have been developed for nearly all bacterial pathogens.76 Rapid and sensitive DNA-based assays that are applicable for the direct detection of enterococci in human clinical 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 clinical samples can come across several problems, such as PCR inhibition by sample-associated components, crosscontamination in sensitive PCR assays, and 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). Although PCR amplification of species-specific targets enables precise identification of known microbial species with short reporting times, it is also important 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 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.77 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,78 multiplex PCR,79– 81 species-specific PCR,82–87 real-time PCR,76,88,89 direct detection of bacteria using fluorescent in situ hybridization (FISH) probes,90–92 and amplification of 16S and 23S ribosomal gene sequences for hybridization to a microarray.93,94 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.

234

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 signals 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.76,88,89,95,96 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 rRNA) or specific genes.91–93 This method is rapid and easy-to-perform and allows the in situ observation of the abundance, spatial distribution, and morphology of bacterial cells.92 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.76 By 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 microarray and laser-scanner apparatus. When applying microarrays for microorganism detection, PCR amplification of target genes is often involved in order to enhance detection sensitivity.76,97–99 Microarray methodology has already been applied for the detection of enterococci in environments such as wastewater76 and artificially contaminated milk samples.94 A summary of available probes for the most relevant enterococcal human pathogens that can be used in qPCR, RT–qPCR, hybridization assays, FISH, and microarrays is presented in Table 22.1. As stated before, a variety of molecular methods have been developed for enterococcal detection and identification at the genus or species level. The detected DNA targets include genes encoding for rRNA,76,84,88,89,103,104 the alpha subunit of RNA polymerase and the phenylalanyl-tRNA synthase,106 the elongation factor EF-Tu,83 the d-alanine:dalanine ligase,80 a penicillin-binding protein,77 the muramidase,87 the alpha subunit of the bacterial ATP synthase,107 the manganese-dependent superoxide dismutase,82,108 and the heat-shock proteins (groESL),76,86,109,110 as well as genus or species-specific antimicrobial determinants.80,81,89 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 22.2. Regarding specific primers for identification at species level, Table 22.2 only includes the most relevant human pathogens (E. faecalis, E. faecium,

Molecular Detection of Human Bacterial Pathogens

E. casseliflavus/E. flavescens, and E. gallinarum) and for information about other enterococcal species, see SemedoLemsaddek et al.140 Ribosomal RNA. rRNA is a universal component of bacterial ribosomes and 5S, 16S, and 23S rRNAs are present at the small (30S) or 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, 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.121 Small subunit ribosomal RNA (16S rRNA) gene sequences are widely used as a 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 intraspecies level, it has been less frequently used, mainly because of 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 rRNA intergenic noncoding region (ITS) may be necessary to distinguish very similar microorganisms. Because of 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 products with complex mixtures of bacteria, making specificity analysis fundamental for validation of the method. Nowadays, several databases compiling full and partial rRNA sequences are available online and updated regularly, allowing fast and easy comparison of data from unknown organisms. These databases include the Ribosomal Database Project II Release 10 (update 14, released on August 31, 2009) that consists of 1074,075 aligned and annotated 16S rRNA sequences, along with seven online analysis tools (http://rdp. cme.msu.edu/). 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, Naser106 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 maximum of 97% rpoA and 86% pheS sequence similarity, pointing to the usefulness of this gene as a target for ­species-level

235

Enterococcus

TABLE 22.1 Probes for Detection and Identification of Enterococci Organism

Target

Eubacteria

16S rRNA 16S rRNA 16S rRNA 16S rRNA

EUB338- GCTGCCTCCCGTAGGAGT GTACAAGGCCCGGGAACGTATTCACC GACATAAGGGGCATGATGATTTGACGT Encl 31- CCCCTTCTGATGGGCAGG

FISH Microarrays Microarrays Microarrays

100 101 101 96

16S rRNAa 16S rRNA

Encl 45- GGGATAACACTTGCAAAC Encl 259- GAAGTCGCGAGGCTAAGC

Microarrays Microarrays

96 96

16S rRNA

ENC221- CACCGCGGGTCCATCCATCA

FISH

92

16S rRNA

ENF191- GAAAGCGCCTTTCACTCTTATGC

FISH

92

23S rRNA

TGGTTCTCTCCGAAATAGCTTTAGGGCTA

TaqMan PCR (qPCR)

88

23S rRNA

GPL813TQb- TGGTTCTCTCCGAAATAGCTTTAGGGCTA

TaqMan PCR (qPCR)

88

vanA enterococci

23S rRNA 23S rRNA vanA gene

Enc01aV- AGGTTAAGTGAATAAGGG ENC176- CAGTTCTCTGCGTCTACCTC vana3- CAACTAACGCGGCACTGTTTCCCAAT

Microarrays FISH TaqMan PCR (qPCR)

96 92 89

vanC enterococci

16S rRNA

EGAC183- CAACTTTCTTCCATGCGGAAAAT

FISH

92

E. casseliflavus

23S rRNA

Eca58- AGCTTGTCCGTACAGGTA

Microarrays

94

E. faecalis

16S rRNA

EFA1- TGATTTGAAAGGCGCTTTCGGGTGTCGCTGATGGATGGAC Microarrays

84

16S rRNA

EFA2- GAAGAACAAGGATGAGAGTAACTGTTCATCCCTTGACGG

Microarrays

84

16S rRNA

EFA3- GAAGTACAACGAGTTGCGAAGTCGCGAGGCTAAGCTAAT Microarrays

84

16S rRNA

CAATTGGAAAGAGGAGTGGCGGACG

TaqMan PCR (qPCR) 102

16S rRNA

TGCCGCATGGCATAAGAGTGAAAGGCGCTTTCGGGTGTC

Microarrays

103

16S rRNA

CAAGGACGTTAGTAACTGAACGTCCCCTGACGGTATCTAA

Microarrays

103

16S rRNA

GAAGTACAACGAGTCGTTAGACCGCGAGGTCATGCAAATC

Microarrays

103

16S rRNA

CCTCGCGGTCTAGCAGCTCGTTGTGCTT

Microarrays

101

16S rRNA

Enfl84- UGCACUCAAUUGGAAAGAGG

FISH

Gram-positive bacteria Enterococcus spp.

Sequence (5′–3′)

Method

Reference

91

16S rRNA

efs129- CCCTCTGATGGGTAGGTT

Microarrays

ITS region

Efs1- TATTTATTGATTAACCTTCT(T)

Hybridization

104

ITS region

Efs2- AAGAAGTGATCAAGACCCA(T)

Hybridization

104

23S rRNA

Efs18i- CGAAATGCTAACAACACC

Microarrays

modified by 93

23S rRNA

Efa54- CAAAAACAACTGGTACAG

Microarrays

96

16S rRNA

TGATTTGAAAGGCGCTTTCGGGTGTCGCTGATGGATGGAC

Microarrays

103

16S rRNA

GAAGAACAAGGATGAGAGTAACTGTTCATCCCTTGACGG

Microarrays

103

16S rRNA

GAAGTACAACGAGTTGCGAAGTCGCGAGGCTAAGCTAAT

Microarrays

103

ITS region

Efm1- TTTTATGAGACGATCGAT(T)

Hybridization

104

ITS region

Efm2- TCTTGATCTAACTTCTAT(T)

Hybridization

104

23S rRNA

Efm09i- GGATGTTACGATTGTGTG

Microarrays

modified by 93

23S rRNA

Efm09- CACACAATCGTAACATCC

Microarrays

96

23S rRNA

Efi58- TGACTCCTCTCCAGACTT

Microarrays

96

23S rRNA

ATCATACGATCAGCCGCAGTGAATA

RT–qPCR

76

23S rRNA

ENU140- TTCACACAATCGTAACATCCTA

FISH

92

23S rRNA

ENU1470- GACTCCTTCAGACTTACTGCTTGG

FISH

92

E. flavescens

23S rRNA

Efl58i- TTCTACCTATACGGACAA

Microarrays

E. gallinarum

ITS region

Egal- GAGTGGACAAGTTAAAGA(T)

Hybridization

23S rRNA

Ega09- CACAACTGTGTAACATCC

Microarrays

96

23S rRNA

EGA141- ATTCACAACTGTGTAACATCCTAT

FISH

92

23S rRNA

Eacdfg57- AGACATATCCATCAGTCT

Microarrays

96

ITS region

Ecasc- GAGTTGAAATGTTAAAAGAG(T)

Hybridization

23S rRNA

Ecafl09i- GGATGTTACGTCTGCGTG

Microarrays

E. faecium

E. flavescens/ E. gallinarum E. casseliflavus/ E. flavescens

96

96 104

104 96 continued

236

Molecular Detection of Human Bacterial Pathogens

TABLE 22.1 (Continued) Probes for Detection and Identification of Enterococci Organism

Target

Sequence (5′–3′)

Method

Reference

E. casseliflavus/ E. gallinarum/ E. flavescens E. faecium

16S rRNA

CGF- CCGTATAACACTATTTTCCGC

Hybridization

105

23S rRNA

Edfm57- CTGCTTGGACAGACATTT

Microarrays

E. faecium

16S rRNA

FMDUR- CATTCAGTTGGGCACTCTAGCAAGA

Hybridization

E. faecium

16S rRNA

Enfm93- CCGGAAAAAGAGGAGUGGC

FISH

91

E. gallinarum/ E. casseliflavus/ E. flavescens E. faecium

16S rRNA

Ecg191- GCGCCTTTCAACTTTCTT

Microarrays

96

16S rRNA

Enc93- GCCACTCCTCTTTTTCCG

Microarrays

96

E. casseliflavus/ E. flavescens

16S rRNA

Ecf459- GGGATGAACATTTTACTC

Microarrays

96

96 105

  Not E. faecalis.

a

TABLE 22.2 Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′)

Product size (bp)

Observations

Reference

Eubacteria 16S rRNA For- GGATTAGATACCCTGGTAGTCC

320

111 in 112

Rev- TCGTTGCGGGACTTAACCCAAC Sm785 For- GGATTAGATACCCTGGTAGTC

~1500

For- GCTGGATCACCTCCTTTC

676/578

Sm785 For + 422Rev

113 114

Uni331 For- TCCTACGGGAGGCAGCAGT

466

115

1554

84, 103

170

79

1522

86

996

116

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 23S rRNA 422 Rev- GGAGTATTTAGCCTT

~1500

Sm785 For + 422Rev

113

L189 Rev- GGTACTTABATGTTTCAGTTC

~1000

ENFE + L189

117

ECST784 For- AGAAATTCCAAACGAACTTG

92

118

900

76

ENC854 Rev- CAGTGCTCTACCTCCATCATT For- AGGAKGTTGGCTTAGAAGCAG Rev- CGCTACCTTAGGACCGTTATAGTTAC rrn 13B For- GTGAATACGTTCCCGGGCCT

~600

16S rRNA + 23S rRNA + ITS

104

6 Rev-GGGTTYCCCCRTTCRGAAAT Chaperonin 60 H279A For- GAIIIIGCIGGIGA(TC)GGIACIACIAC

600

109

H280A Rev- (TC)(TG)I(TC)(TG)ITCICC(AG)AAICCIGGIGC(TC)TT continued

237

Enterococcus

TABLE 22.2 (Continued) Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′) 590 For- GGNGACGGNACNACNACNGCAACNGT 590 Rev- TCNCCRAANCCNGGYGCNTTNACNGC

Product Size (bp)

Observations

Reference

589

110

803

83

Elongation factor EF-Tu (tuf) U1 For- AAYATGATIACIGGIGCIGCICARATGGA U2 Rev- AYRTTITCICCIGGCATIACCAT Lactic acid bacteria α subunit of ATP synthase (atpA) atpA-20 For- TAYRTYGGKGAYGGDATYGC

1102

107

494

106

431

106

815

106

atpA-27 Rev- CCRCGRTTHARYTTHGCYTG Phenylalanyl-tRNA synthase (pheS) pheS-21 For- CAYCCNGCHCGYGAYATGC pheS-22 Rev- CCWARVCCRAARGCAAARCC pheS-21 For- CAYCCNGCHCGYGAYATGC pheS-23 Rev- GGRTGRACCATVCCNGCHCC RNA polymerase a subunit (rpoA) rpoA-21 For- ATGATYGARTTTGAAAAACC rpoA-23 Rev- ACHGTRTTRATDCCDGCRCG Genus Enterococcus 16S rRNA 733

119

115

120

337

121

92

88

320 + 420

122,123 in 124

1609

110

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

112

83

Manganese-dependent superoxide dismutase (sodA) D1 For- CCITAYICITAYGAYGCIYTIGARCC D2 Rev- ARRTARTAIGCRTGYTCCCAIACRTC

480

82

E1 For- TCAACCGGGGAGGGT E2 Rev- ATTACTAGCGATTCCGG 16S rRNA Ec-ssu1 For- GGATAACACTTGGAAACAGG Ec-ssu1 Rev- TCCTTGTTCTTCTCTAACAA 16S rRNA For- ATCAGAGGGGGATAACACTT Rev- ACTCTCATCCTTGTTCTTCTC 23S rRNA ECST784 For - AGAAATTCCAAACGAACTTG ENC854 Rev- CAGTGCTCTACCTCCATCATT ITS region For- CAAGGCATCCACCGT Rev- GAAGTCGTAACAAGG Chaperonin 60 GroES EntGroES For- TTAAAACCATTAGGCGATCG EntGroES Rev- CCCATNCCCATNGANGGRTCCAT

E. faecalis 16S rRNA Efs130 For- AACCTACCCATCAGAGGG Efs490 Rev- GACGTTCAGTTACTAACG For- CGCTTCTTTCCTCCCGAGT Rev- GCCATGCGGCATAAACTG

360 143

125 126 102 continued

238

Molecular Detection of Human Bacterial Pathogens

TABLE 22.2 (Continued) Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′)

Product Size (bp)

For- TACTGACAAACCATTCATGATG

Observations

Reference

112

  127

138

79

Rev- AACTTCGTCACCAACGCGAAC 72 For- CCGAGTGCTTGCACTCAATTGG 210 Rev- CTCTTATGCCATGCGGCATAAAC ENFE For- GTCGCTAGACCGCGAGGTCATGA

~1000

ENFE + L189

128

Chaperonin 60 GroESL 185

110

64

76

650

86

941

80

475

129

803

86

360

108

444

77

518

130

EE1 For- TGTGGTATCGGAGCTGCAG

430

129

EE2 Rev- ATAGTTTAGCTGGTAAC For- TGTGGTATCGGAGCTGCAG Rev- GTCGATTCTCGCTAATCC

513

131

810

132

941

129

EfGroES For- GGAATTGTTCTTGCATCCGT EfGroES Rev- ACAATTAAGTATTCTACGCC For- TGTGGCAACAGGGATCAAGA Rev- TTCAGCGATTTGACGGATTG Efes For- GTGTTAAAACCATTAGGCGAT Efes Rev- AAGCCTTCACGAACAATGG d-Alanine/d-alanine

ligase (ddl)

For- ATCAAGTACAGTTAGTCTTTATTAG Rev- ACGATTCAAAGCTAACTGAATCAGT DD13 For- CACCTGAAGAAACAGGC DD13 Rev- ATGGCTACTTCAATTTCACG Iron sulfur–binding protein Efis For- ATGCCGACATTGAAAGAAAAAATT Efis Rev- TCAATCTTTGGTTCCATCTCT Manganese-dependent superoxide dismutase (sodA) FL1 For- ACTTATGTGACTAACTTAACC FL2 Rev- TAATGGTGAATCTTGGTTTGG Penicillin-binding protein (pbp5) For- CATGCGCAATTAATCGG Rev- CATAGCCTGTCGCAAAAC Ef0027- unknown function For- GCCACTATTTCTCGGACAGC Rev- GTCGTCCCTTTGGCAAATAA vane

vanG For- CGGTTGTGCCGTACTTGGC Rev- GGGTAAAGCCATAGTCTGGGGC EG1 For- CGGCATCCGCTGTTTTTGA EG2 Rev- GAACGATAGACCAATGCCTT

E. faecalis and E. faecium vanA vana3 For- CTGTGAGGTCGGTTGTGCG vana3 Rev- TTTGGTCCACCTCGCCA

64

89

vanB For- GTGACAAACCGGAGGCGAGGA Rev- CCGCCATCCTCCTGCAAAAAA

433

133

For- CATCGCCGTCCCCGAATTTCAAA

298

81

Rev- GATGCGGAAGATACCGTGGCT continued

239

Enterococcus

TABLE 22.2 (Continued) Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′)

Product Size (bp)

EB3 For- ACGGAATGGGAAGCCGA

Observations

647

Reference 129

EB4 Rev- TGCACCCGATTTCGTTC For- CATCGCCGTCCCCGAATTTCAAA

297

134

500

129

630

135

461

136

628

137

Rev- GATGCGGAAGATACCGTGGCT vanD ED1 For- TGTGGGATGCGATATTCAA ED2 Rev- TGCAGCCAAGTATCCGGTAA For- GARGATGGITSCATMCARGGW Rev- MGTRAAICCIGGCAKRGTRTT For-TAAGGCGCTTGCATATACCG Rev- TGCAGCCAAGTATCCGGTAA vanD-U1 TATTGGAATCACAAAATCCGG vanD-U2 CGGCTGTGCTTCCTGATG E. faecium 23S rRNA Ef Rev- CACACAATCGTAACATCCTA d-Alanine/d-alanine

676/578

95

550

80

1091

129

215

108

658

138

ligase (ddl)

For- GCAAGGCTTCTTAGAGA Rev- CATCGTGTAAGCTAACTTC FAC1 For- GAGTAAATCACTGAACGA FAC2 Rev- CGCTGATGGTATCGATTCAT Manganese-dependent superoxide dismutase (sodA) FM1 For- GAAAAAACAATAGAAGAATTAT FM2 Rev- TGCTTTTTTGAATTCTTCTTTA Unknown region For- TTGAGGCAGACCAGATTGACG Rev- TATGACAGCGACTCCGATTCC

Other enterococcal species Manganese-dependent superoxide dismutase (sodA) 288

E. casseliflavus

108

FV1 For- GAATTAGGTGAAAAAAAAGTT

284

E. flavescens

108

FV2 Rev- GCTAGTTTACCGTCTTTAACG GA1 For- TTACTTGCTGATTTTGATTCG GA2 Rev- TGAATTCTTCTTTGAAATCAG

173

E. gallinarum

108

1030

Several enterococcal species

133

377

Several enterococcal species

81

783

Several enterococcal species

134

822

E. gallinarum and E. casseliflavus/E. flavescens

80

438

E. gallinarum and E. casseliflavus/E. flavescens

81

CA1 For- TCCTGAATTAGGTGAAAAAAC CA2 Rev- GCTAGTTTACCGTCTTTAACG

vanA For- CATGAATAGAATAAAAGTTGCAATA Rev- CCCCTTTAACGCTAATACGATCAA For- TCTGCAATAGAGATAGCCGC Rev- GGAGTAGCTATCCCAGCATT For- GCTATTCAG CTGTACTC Rev- CAGCGGCCATCATACGG vanC1 For- GGTATCAAGGAAACCTC Rev- CTTCCGCCATCATAGCT For- GACCCGCTGAAATATGAAG Rev- CGGCTTGATAAAGATCGGG continued

240

Molecular Detection of Human Bacterial Pathogens

TABLE 22.2 (Continued) Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′) For- GGTATCAAGGAAACCTC

Product Size (bp)

Observations

Reference

822

E. gallinarum and E. casseliflavus/E. flavescens

134

815/827

E. gallinarum and E. casseliflavus/E. flavescens

129

430

E. gallinarum and E. casseliflavus/E. flavescens

81

484

E. gallinarum and E. casseliflavus/E. flavescens

139

439

E. gallinarum and E. casseliflavus/E. flavescens

134

Rev- CTTCCGCCATCATAGCT vanC1/C2 EC5 For- ATGGATTGGTAYTKGTAT EC8 Rev- TAGCGGGAGTGMCYMGTAA vanC2 For- CTCCTACGATTCTCTTG Rev- CGAGCAAGACCTTTAAG vanC2/C3 For-CGGGGAAGATGGCAGTAT Rev- CGCAGGGACGGTGATTTT For- CTCCTACGATTCTCTTG Rev- CGAGCAAGACCTTTAAG

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 genome83,141 making this region a good target for diagnostic purposes. Ke and coworker83 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 coworkers80 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 speciesspecific PCR of E. durans and E. hirae targeting the ddl gene were also developed and validated by Knijff and coworkers.142 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 speciesspecific probe143 and was subsequently used in competitive PCR for quantification of nonculturable E. faecalis cells.77 Muramidase. In 2006, Arias and coworkers87 searched the GenBank for sequences to be applied for the identification of E. hirae. During this process, an E. durans muramidase gene (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.144 The aim of the study developed by Naser and coworkers107 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 speciesspecific primers have been described to demonstrate this potential. Manganese-Dependent Superoxide Dismutase. In 2000, Poyart and coworkers82 sequenced the manganesedependent 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 primers108 that can be applied to identify 23 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.145 Goh and coworkers109 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 coworkers110 determined the groESL gene’s full-length sequence of E. faecalis and used this information to develop primers specific for this species. Antimicrobial Resistance. Over the years, several authors used multiplex PCR to both detect the presence of vancomycin

241

Enterococcus

resistance determinants and identify enterococci to the species level.80,112,129,139,146 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 teicoplanin147 and have been found in several enterococcal species.148 The vanB genes confer resistance to various concentrations of vancomycin but not teicoplanin149; 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;136,150 vanD determinants are constitutively expressed in E. faecalis but inducible in E. faecium.136 The vanE gene is induced by low levels of vancomycin131 and has only been detected in one strain of E. faecalis. The vanC1 and vanC2/C3 genes are specific to the motile VRE species E. gallinarum and E. casseliflavus/E. flavescens, respectively.80 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.132

22.2  METHODS 22.2.1  Sample Preparation It is often difficult to establish whether an Enterococcus strain is contributing to the infection or whether it is colonizing a human or an animal. Bacteremia as major cause of nosocomial infection represents an important source of morbidity and mortality. Gram-positive microorganisms, especially Staphylococcus and E. faecalis, account for the majority of episodes of bacteremia in critically ill patients in intensive care units (ICU). The rapid and reliable detection and subsequent identification of microorganisms, namely, Enterococcus, from blood is critical so that appropriate antimicrobial therapy can be provided. The same principle has to apply to the rapid and reliable detection of microbial samples from stool samples, oral samples, urine, pus, or exudates. The appropriate collection and transport of specimens are critical to ensure that proper diagnostic information is given by the microbiology laboratory.151,152 Also the initial processing of clinical specimens, which is a multifaceted endeavor as one has to consider specimen type and its anatomical origin, is a critical step. Description of the pretreatment and treatment of the specimens received in laboratory for aerobic microbiology in which Enterococcus is included have been gathered by York in 2007,153 but generally they start by microscopical observation, Gram stain and isolation from the clinical samples in liquid and solid media, followed by identification procedures. Selective agents that are commonly used in selective media to isolate enterococci include: sodium azide, thallous acetate, kanamycin, and gentamicin. Other limiting ingredients or factors are crystal violet, Tween

80, carbonate, and bile salts. Special growth conditions may be also used, like low pH (e.g., 6.0 or 6.2) or elevated incubation temperature (42°C or 45°C). Nutritional requirements vary among the members of the genus, but it usually an advantage to use peptones, beef or yeast extract. Considering the mentioned factors it is easy to understand why there are so many varieties of selective media and discrepant results in the literature. ABA (esculin bile azide agar), KAA (kanamycin aesculin azide agar), CATC (citrate azide tween carbonate agar), ME (m-Enterococcus agar), TITG (thallous acetate tetrazolium glucose agar), and KA (crystal violet azide agar) are commonly used to cultivate enterocci.154–156 Laboratories vary on the media they use depending on the type of sample; for instance, some use KAA agar to isolate enterococci from feces and cysteine-lactose-electrolyte-deficient (CLED) agar can be used in bacteriological studies of urine.157 The detection and identification of vancomycin-resistant enterococci (VRE) led to the development of special media and procedures to detect and identify them. Some authors demonstrated that VRE grew on Mueller-Hinton agar with 6–12 µg/mL of vancomycin, although 6 µg/mL of vancomycin seems to be the optimum concentration.12,158 Chromogenic media are now also widely used namely for cell counts and identification of enterococci from urine or stool samples, either if they are vancomycin-resistant or not. Media like BBL CHROMagar (BD Diagnostics), Chromogenic VRE (AES Laboratoire), chromID VRE (bioMérieux), VRE agar (Oxoid) or even bile esculin agar supplemented with 6 µg/mL of vancomycin (BEV), and bile esculin azide vancomycin (BEAV) agar have been used.159–162 After isolation of potential Enterococcus strains from clinical samples, using any of the media already mentioned, strains can be grown in trypticase-soy-5% sheep blood agar, brain heart infusion-5% sheep blood agar, or any blood agar base containing 5% animal blood, as well as brain heart infusion (BHI), MRS agar or broth (DeMan, Rogosa, Sharpe) or M17agar or broth to be further characterized or identified by PCR. All enterococci grow at 35ºC–37ºC and do not require an atmosphere containing increased levels of carbon dioxide.

22.2.2  Detection Procedures 22.2.2.1  Semiautomatic and Automatic Identification The blood culture is the gold standard for diagnosing bacteria, but it may take more than two days for results to be available. Semiautomatic or automatic methods to test blood culture or other body fluids and to identify enterococci directly from samples, like BACTEC Peds PLUS/F and BACTEC 9240 that use spectrophotometry to monitor the CO2 being produced during growth, are now commonly used, and the results have been compared to other systems of identification.163,164 Other systems used for the identification of clinically isolated enterococci are kits like API 20 Strep and API Rapid ID 32 STREP (bioMérieux) and BBL Crystal gram-positive and BBL Crystal Rapid gram-positive (BD Diagnostics), that were already compared in terms of

242

Molecular Detection of Human Bacterial Pathogens

specificity.165 There are also systems that identify and at the same time test the antibiotic susceptibility of the strains like the Vitek 2 and Vitek Compact Systems (bioMerieux). Also very used are the identification systems Bact/ALERT 3D (bioMérieux), BACTEC 9000 (BD Diagnostics), Versa Trek (Trek), or Phoenix 100 (Becton Dickson).165–168 22.2.2.2  M  ultiplex PCR for the Detection of Enterococcus Species Although most of the routine identification of pathogens in clinical samples start with bacterial isolation, the advent of molecular approaches opens the road for direct application of such methods without culturing. In this context, the experimental procedure hereby described may be applied using DNA extracted both from isolated strains and clinical samples. A list of kits that can be used to extract DNA, directly from biological samples or from isolated colonies, has already been compiled.140 After DNA extraction, it is proposed that the detection/ identification of Enterococcus can follow the steps shown in Clinical sample

DNA extraction

Primers

E1 + E2 FL1 + FL2 FM1 + FM2

Target species

Identification by genus- and species-specific multiplex PCR

E. faecalis 360 bp

E. casseliflavus 288 bp

E. faecium 215 bp

E. gallinarum 173 bp

E1 + E2 CA1 + CA2 GA1 + GA2

Screening of putative virulence factors by multiplex PCR

A1 + A2 B1 + B2 C1 + C2 D1 + D2

CYT I + CYT IIb ESP 14F + ESP 12R ASA 11 + ASA 12 GEL 11 + GEL 12 HYL n1 + HYL n2

Target genes

Primers

Screening of antimicrobial resistance by multiplex PCR

vanA vanB vanC1 vanC2/C3

cytolysin -cylAenterococcal surface protein -espaggregation substance -asalgelatinase -gelEhialuronidase -hyl-

FIGURE 22.1  Flowchart of the multiplex-PCR-based procedures for the detection and/or identification of enterococci at genus and species level, as well as for the assessment of antimicrobial and virulence determinants.

the flow chart of Figure 22.1. The procedure is a genus- and species-specific multiplex PCR, adapted from the 23-species detection methodology of Jackson and coworkers,108 enabling the detection of E. faecalis, E. faecium, E. casseliflavus, and E. gallinarum. Because this approach combines the genus-specific primers (E1/E2) described by Deasy and coworkers119 with primers for the superoxide dismutase (sod) gene108 specific for E. faecalis (FL1/FL2), E. faecium (FM1/FM2), E. casseliflavus (CA1/ CA2), and E. gallinarum (GA1/GA), it provides a rapid and reliable method for the detection and/or identification of the most important human pathogenic enterococcal species. Amplification procedure consists of the preparation of a PCR master mix (60 µL) containing 3 mM MgCl2, 0.2 mM dNTP mix, 3.5 U of high fidelity DNA polymerase, and 3.75  μL (16 μM) of genus-specific primer set (E1/E2). The master mix should be equally divided in three separated vials in order to use the primer combinations depicted in Figure 22.1 (for sequence see Table 22.2) and have a positive control of the reaction. To vial number 1 add 2.5 µL (16 µM) of primers FL1/FL2 and 1.25 μL (16 μM) of primers FM1/FM2; to vial number 2 add 2.5 µL (16 µM) of primers GA1/GA2 and 1.25 μL (16 μM) of primers CA1/CA2; to vial number 3 (positive control) add DNA from a nonpathogenic strain of Enterococcus. PCR reactions are performed in a final volume of 22.5 μL after the addition to each vial of 2.5 μL of test DNA, extracted either from clinical sample or isolated strain. The PCR amplification program includes an initial denaturation at 95°C for 4 min; 30 cycles of 95°C for 30 s, 55°C for 1 min, and 72°C for 1 min; a final 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 used as standard. Following electrophoresis, the agarose gels are visualized and photographed under UV light. Since the presence of enterococcal isolates carrying virulence and antimicrobial resistance determinants turns the pathogenicity potential of these bacteria higher, we suggest an integrative approach leading to species identification and assessment for the presence of vancomycin resistance and virulence determinants (Figure 22.1). Although this approach is not essential for treating enterococcal infections, it may contribute to the implementation of more adequate antimicrobial treatment and/or to the implementation of measures aimed at the elimination of more virulent enterococci from hospital settings. Screening for the presence of antimicrobial resistance and virulence determinants can also be performed by multiplex PCR, using the methodologies described by DutkaMalen et al.80 and Vankerckhoven et al.,169 respectively. 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 22.2 and Figure 22.1). 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 clinical

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Enterococcus

samples). When only the genus-specific amplicon is visualized, the presence of another enterococcal species is inferred. If no amplicons are observed, with the exception of the positive control reaction tube (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).

22.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES Enterococcus are generally not as virulent as other grampositive bacteria but can occur as a polymicrobial infection in debilitated hosts.170 Resistance of enteroccci to multiple antimicrobial agents, including vancomycin, can occur in health care centers. Considering that the most problematic enteroccci are VREs, the infection control has to pass through careful analysis on VRE spread. Since colonized patients are the primary reservoir of VRE, a patient-to-patient transmission occurs in the first place. As hospital staff and environmental contamination, through medical devices or surroundings of colonized patients, also plays a critical role in spread, all these aspects have to be taken into consideration.170 The Hospital Infection Control Practices Advisory Committee (HIPAC) has published in 1995 “Recommendations for Preventing the Spread of Vancomycin Resistance” (Centers for Disease Control and Prevention. Recommendations for Preventing the Spread of Vancomycin Resistance).171 These measures include the appropriate use of vancomycin, the establishment of education programs, the clear understanding of the role of microbiology laboratory, the prevention and control of nosocomial transmission of VRE, and the isolation precautions to prevent patient-to-patient transmission. More recently, research on antimicrobial stewardship, meaning the optimal selection dose and duration of an antimicrobial that results in best clinical outcome for the treatment and prevention of infection, with minimum toxicity to the patient and minimal impact on subsequent resistance and development, has produced more and more data.172–174 Health care facilities should then adopt antimicrobial stewardship programs (ASPs) and infection control programs (ICPs) to monitor antimicrobial use while simultaneously optimizing treatment, outcome, and cost. The IDSA/SHEA guidelines for developing an institutional program to enhance antimicrobial stewardship can be used as a starting point to control the threat.175,176 All these evaluations have to take on board, not only healthy people, who seem to be unaffected by enterococcal strains, but also the elderly population, infants, and immunocompromised people. The microbiological risk assessment of emerging pathogens has to be performed, and global data indicate that the epidemiology of human diseases is changing because of changes in lifestyles and improved surveillance and recovery/detection methods.177,178 Pathogen control, and especially VRE, depends on the laboratory’s ability to detect,

identify, and recognize those agents; therefore, in the case of Enterococcus, a detailed combination of a more specific genus and species method is proposed here based on the work of Jackson and coworkers.108 PCR methods in general, and real-time PCR methods in particular, are increasingly used for the detection and quantification of pathogens in clinical samples. Nonetheless, the new species that are constantly appearing in the literature, calling for new and/or reassessed methods of identification and quantification as well as for the need to always be alert to new antibiotic resistances, are a constant challenge and constitute new routes of needed research in the area of work with Enterococcus.

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 project BIOSAFE SSPS-CT-2006–022725. T. Semedo-Lemsaddek thanks the financial support of the FCT Pos-Doctoral fellowship SFRH/BPD/20892/2004.

REFERENCES





1. Thiercelin, E., and Jouhaud, L., Reproduction de l´entérocoque; taches centrales; granulation périphériques et microblastes, C. R. Soc. Biol., 55, 686, 1903. 2. Murray, B.E., The life and times of the Enterococcus, Clin. Microbiol. Rev., 46, 65, 1990. 3. Kalina, A.P., The taxonomy and nomenclature of enterococci, Int. J. Syst. Bacteriol., 20, 185, 1970. 4. Schleifer, K.H., and Kilpperbalz, R., Transfer of Streptococcus faecalis and Streptococcus faecium to the Genus Enterococcus nom. rev. as Enterococcus faecalis comb. nov. and Enterococcus faecium comb. nov., Int. J. Syst. Bacteriol., 34, 31, 1984. 5. Ludwig, W. et al., The phylogenetic position of Streptococcus and Enterococcus, J. Gen. Microbiol., 131, 543, 1985. 6. Kohler, W., The present state of species within the genera Streptococcus and Enterococcus, Int. J. Med. Microbiol., 297, 133, 2007. 7. Garrity, G.M. et al., The Bacteria: Phylum Firmicutes: Class “Bacilli.” In Taxonomic Outline of the Bacteria and Archea (TOBA), Release 7.7, Part 9, 2007. 8. Devriese, L.A. et al., Identification of aesculin-hydrolyzing streptococci, lactococci, aerococci and enterococci from subclinical intramammary infections in dairy cows, Vet. Microbiol., 70, 87, 1999. 9. Devriese, L.A., Baele, M., and Butaye, P., The genus Enterococcus, In Dworkin, M. et al. (Editors), The Prokaryotes, pp. 163–174, Springer, Heidelberg, 2002. 10. Holt, J.G. et al., Bergey’s Manual of Determinative Bacteriology, 9th ed., William & Wilkins, Baltimore, MD, 1994. 11. Devriese, L.A. et al., Identification of Enterococcus species isolated from foods of animal origin, Int. J. Food Microbiol., 26, 187, 1995.

244 12. Facklam, R.R., Carvalho, M.G.S., and Teixeira L.M., History, taxonomy, biochemical characteristics, and antibiotic susceptibility testing of enterococci. In Gilmore, M.S. et al. (Editors), The Enterococci: Pathogenesis, Molecular Biology and Antibiotic Resistance, pp. 1–46, ASM Press, Washington, D.C., 2002. 13. Doming, K.J., Mayer, H.K., and Kneifel, W., Methods used for the isolation, enumeration, characterization and identification of Enterococcus spp. 2. Pheno- and genotypic criteria, Int. J. Food Microbiol., 88, 165, 2003. 14. Foulquié Moreno, M.R. et al., The role and application of enterococci in food and health, Int. J. Food Microbiol., 106, 1, 2006. 15. Ogier, J.C., and Serror, P., Safety assessment of dairy microorganisms: The Enterococcus genus, Int. J. Food Microbiol., 126, 291, 2008. 16. Kayser, F.H., Safety aspects of enterococci from the medical point of view, Int. J. Food Microbiol., 88, 255, 2003. 17. Lund, B. et al., Frequent transmission of enterococcal strains between mechanically ventilated patients treated at an intensive care unit, J. Clin. Microbiol., 40, 2084, 2002. 18. Low, D.E. et al., Clinical prevalence, antimicrobial susceptibility, and geographic resistance patterns of enterococci: Results from the SENTRY Antimicrobial Surveillance Program, 1997–1999, Clin. Infect. Dis., 32, S133, 2001. 19. Schaberg, D.R., Culver, D.H., and Gaynes, R.P., Major trends in the microbial etiology of nosocomial infection, Am. J. Med., 91, S72, 1991. 20. Murray, B.E., Diversity among multidrug-resistant enterococci, Emerg. Infect. Dis., 4, 37, 1998. 21. Wisplinghoff, H. et al., Nosocomial bloodstream infections in US hospitals: Analysis of 179 cases from a perspective nation wide surveillance study, Clin. Infect. Dis., 39, 309, 2004. 22. Huycke, M.M., Sahm, D.F., and Gilmore, M.S., Multiple drug resistant enterococci: The nature of the problem and an agenda for the future, Emerg. Infect. Dis., 4, 239, 1998. 23. Bouza, E. et al., A European perspective on nosocomial urinary tract infections II. Report on incidence, clinical characteristics and outcome (ESGNI-004 study). European Study Group on Nosocomial Infection, Clin. Microbiol. Infect., 7, 532, 2001. 24. Reynolds, R. et al., Antimicrobial susceptibility of the pathogens of bacteraemia in the UK and Ireland 2001–2002: The BSAC Bacteraemia Resistance Surveillance Programme, J. Antimicrob. Chemother., 53, 1018, 2004. 25. Coque, T.M. et al., Population structure of Enterococcus faecium causing bacteremia in a Spanish University Hospital: Setting the scene for a future increase in vancomycin resistance? Antimicrob. Agents Chemother., 49, 2693, 2005. 26. Giraffa, G., Enterococci from foods, FEMS Microbiol. Rev., 26, 163, 2002. 27. Klein, G., Taxonomy, ecology and antibiotic resistance of enterococci from food and the gastro-intestinal tract, Int. J. Food Microbiol., 88, 123, 2003. 28. Mundy, L.M., Sahm, D.F., and Gilmore, M., Relationships between enterococcal virulence and antimicrobial resistance, Clin. Microbiol. Rev., 13, 513, 2000. 29. Wenzel, R.P., and Edmond, M.B., The impact of hospitalacquired bloodstream infections, Emerg. Infect. Dis., 7, 174, 2001. 30. Uttley, A.H.C. et al., Vancomycin-resistant enterococci, Lancet, 1, 57, 1988. 31. Menichetti, F., Current and emerging serious Gram-positive infections, Clin. Microbiol. Infect., 11, 22, 2005.

Molecular Detection of Human Bacterial Pathogens 32. Klare, I. et al., Enterococcus faecium strains with vanA mediated high-level glycopeptides resistance isolated from animal foodstuffs and fecal samples of humans in the community, Microb. Drug Resist., 1, 265, 1995. 33. Aarestrup, F.M., Occurrence of glycopeptide resistance among Enterococcus faecium isolates from conventional and ecological poultry farms, Microb. Drug Resist., 1, 255, 1995. 34. Bager, F. et al., Avoparcin used as a growth promoter is associated with the occurrence of vancomycin resistant Enterococcus faecium on Danish poultry and pigs farms, Prev. Vet. Med., 31, 95, 1997. 35. Wegener, H.C. et al., Use of antimicrobial growth promoters in food animals and Enterococcus faecium resistance to therapeutic antimicrobial drugs in Europe, Emerg. Infect. Dis., 5, 329, 1999. 36. Vergis, E.N. et al., Determinants of vancomycin resistance and mortality rates in enterococcal bacteremia: A prospective multicenter study, Ann. Intern. Med., 135, 484, 2001. 37. Byers, K.E. et al., A hospital epidemic of vancomycin-resistant Enterococcus: Risk factors and control, Infect. Control Hosp. Epidemiol., 22, 140, 2001. 38. Falk, P.S. et al., Outbreak of vancomycin-resistant enterococci in a burn unit, Infect. Control Hosp. Epidemiol., 21, 575, 2000. 39. Jett, B.D., Huycke, M.M., and Gilmore, M.S., Virulence of enterococci, Clin. Microbiol. Rev., 7, 462, 1994. 40. Shankar, V. et al., Infection derived-Enterococcus faecalis strains are enriched in esp, a gene encoding a novel surface protein, Infect. Immun., 1, 193, 1999. 41. Shankar, N. et al., Role of Enterococcus faecalis surface protein Esp in the pathogenesis of ascending urinary tract infection, Infect. Immun., 69, 4366, 2001. 42. Toledo-Arana, A. et al., The enterococcal surface protein, Esp, is involved in Enterococcus faecalis biofilm formation, Appl. Environ. Microbiol., 67, 4538, 2001. 43. Dunny, G.M., Leonard, B.A.B., and Hedberg, P.J., Pheromone inducible conjugation in Enterococcus faecalis. Interbacterial and host parasite chemical communication, J. Bacteriol., 177, 871, 1995. 44. Wirth, R., The Sex pheromone system of Enterococcus faecalis: More than just a plasmid collection mechanism, Eur. J. Biochem., 222, 235, 1994. 45. Sartingen, S. et al., Aggregation substance increases adherence and internalization, but not translocation, of Enterococcus faecalis through different intestinal epithelial cells in vitro, Infect. Immun., 68, 6044, 2000. 46. Suβmuth, S.D. et al., Aggregation substance promotes adherence, phagocytosis, and intracellular survival of Enterococcus faecalis within human macrophages and suppresses respiratory burst, Infect. Immun., 68, 4900, 2000. 47. Wells, C.L. et al., Inducible expression of Enterococcus faecalis aggregation substance surface protein facilitates bacterial internalization by cultured enterocytes, Inf. Immun., 68, 7190, 2000. 48. Hirt, H., Erlandsen, S.L., and Dunny, G.M., Heterologous inducible expression of Enterococcus faecalis pCF10 aggregation substance Asc10 in Lactococcus lactis and Streptococcus gordonii contributes to cell hydrophobicity and adhesion to fibrin, J. Bacteriol., 182, 2299, 2000. 49. Rakita, R.M. et al., Enterococcus faecalis bearing aggregation substance is resistant to killing by human neutrophils despite phagocytosis and neutrophil activation, Infect. Immun., 67, 6067, 1999. 50. Schlievert, P.M. et al., Aggregation and binding substances enhance pathogenicity in rabbit models of Enterococcus faecalis endocarditis, Infect. Immun., 66, 218, 1998.

Enterococcus 51. Jett, B.D., Atkuri, R.V., and Gilmore, M.S., Enterococcus faecalis localization in experimental endophthalmitis: Role of plasmid-encoded aggregation substance, Infect. Immun., 66, 843, 1998. 52. Lowe, A.M., Lambert, P.A., and Smith, A.W., Cloning of an Enterococcus faecalis endocarditis antigen homology with adhesins from some oral streptococci, Infect. Immun., 63, 703, 1995. 53. Singh, K.V. et al., In vivo testing of an Enterococcus faecalis efaA mutant and use of efaA homologs for species identification, FEMS Immunol. Med. Microbiol., 21, 323, 1998. 54. Haas, W., and Gilmore, M.S., Molecular nature of a novel bacterial toxin: The cytolysin of Enterococcus faecalis, Med. Microbiol. Immun., 187, 183, 1999. 55. Ike, Y., Hashimoto, H., and Clewell, D.B. Hemolysin of Streptococcus faecalis subspecies zymogenes contributes to virulence in mice, Infect. Immun., 45, 528, 1994. 56. Johnson, A.P., The Pathogenicity of enterococci, J. Antimicrob. Chemoth., 33, 1083, 1994. 57. Coque, T.M. et al., Incidence of hemolysin, gelatinase, and aggregation substance among enterococci isolated from patients with endocarditis and other infections and from feces of hospitalized and community based persons, J. Infect. Dis., 171, 1223, 1995. 58. Nakayama, J., Kariyama, R., and Kumon, H., Description of a 23.9-kilobase chromosomal deletion containing a region encoding fsr genes which mainly determines the gelatinase negative phenotype of clinical isolates of Enterococcus faecalis in urine, Appl. Environ. Microbiol., 68, 3152, 2002. 59. Pillai, S.K. et al., Prevalence of the fsr locus in Enterococcus faecalis infection, J. Clin. Microbiol., 40, 2651, 2002. 60. Qin, X. et al., Effects of Enterococcus faecalis fsr genes on production of gelatinase and a serine protease and virulence, Infect. Immun., 68, 2579, 2000. 61. Willems, R.J.L. et al., Variant esp gene as a marker of a distinct genetic lineage of vancomycin-resistant Enterococcus faecium spreading in hospitals, Lancet, 357, 853, 2001. 62. Rice, L.B. et al., A potential virulence gene, hyl (Efm), predominates in Enterococcus faecium of clinical origin, J. Infect. Dis., 2003, 187, 508, 2003. 63. Hacker, J., and Kaper, J.B., Pathogenicity islands and the evolution of microbes, Annu. Rev. Microbiol., 54, 641, 2000. 64. Shankar, N., Baghdayan, A.S., and Gilmore, M.S., Modulation of virulence within a pathogenicity island in vancomycinresistant Enterococcus faecalis, Nature, 417, 746, 2002. 65. Leavis, H. et al., A novel putative enterococcal pathogenicity island linked to the esp virulence gene of Enterococcus faecium and associated with epidemicity, J. Bacteriol, 186,672, 2004. 66. Nallapareddy, S.R. et al., Molecular characterization of a widespread, pathogenic, and antibiotic resistance-receptive Enterococcus faecalis lineage and dissemination of its putative pathogenicity island, J. Bacteriol., 187, 5709, 2005. 67. Rahman, I. et al., Methionine uptake and cytopathogenicity of viable but nonculturable Shigella dysenteriae type 1, Appl. Environ. Microbiol., 60, 3573, 1994. 68. Lleo, M.D., Tafi, M.C., and Canepari, P., Nonculturable Enterococcus faecalis cells are metabolically active and capable of resuming active growth, Syst. Appl. Microbiol., 21, 333, 1998. 69. Colwell, R.R., and Huq, A., Vibrios in the environment: Viable but nonculturable Vibrio cholerae. In Kaye, T. (Editor), Vibrio and Cholera: Molecular Global Perspectives, pp. 117–133, ASM Press, Washington D.C., 1994.

245 70. Pruzzo, C. et al., In vitro adhesion to human cells by viable but nonculturable Enterococcus faecalis, Curr. Microbiol., 45, 105, 2002. 71. Lleo, M.D. et al., Vancomycin resistance is maintained in enterococci in the viable but nonculturable state and after division is resumed, Antimicrob. Agents Chemother., 47, 1154, 2003. 72. Lleo, M.D. et al., mRNA detection by reverse transcription-PCR for monitoring viability over time in an Enterococcus faecalis viable but nonculturable population maintained in a laboratory microcosm, Appl. Environ. Microbiol., 66, 4564, 2000. 73. Arana, I. et al., Detection and enumeration of viable but nonculturable transconjugants of Escherichia coli during the survival of recipient cells in river water, J. Appl. Microbiol., 83, 340, 1997. 74. Oliver, J.D., Formation of viable but nonculturable cells. In Kjelleberg, S. (Editor), Starvation in Bacteria, pp. 239–272, Plenum Press, New York, 1993. 75. Lleo, M.M. et al., Resuscitation rate in different enterococcal species in the viable but non-culturable state, J. Appl. Microbiol., 91, 1095, 2001. 76. Lee, D.Y., Shannon, K., and Beaudette, L.A., Detection of bacterial pathogens in municipal wastewater using an oligonucleotide microarray and real-time quantitative PCR, J. Microbiol. Methods, 65, 453, 2006. 77. Lleo, M.D. et al., Competitive polymerase chain reaction for quantification of nonculturable Enterococcus faecalis cells in lake water, FEMS Microbiol. Ecol., 30,345, 1999. 78. Monstein, H.J. et al., Division of the genus Enterococcus into species groups using PCR-based molecular typing methods, Microbiology, 144, 1171, 1998. 79. Sedgley, C.M. et al., Quantitative real-time PCR detection of oral Enterococcus faecalis in humans, Arch. Oral Biol., 50, 575, 2005. 80. Dutka-Malen, S., Evers, S., and Courvalin, P., Detection of glycopeptideresistance genotypes and identification to the species level of clinically relevant enterococci by PCR, J. Clin. Microbiol., 33, 1434, 1995. 81. Klare, I., and Witte, W., Glycopeptide resistant Enterococci— origin, spread, transferable resistance and significance, Wien. Klin. Wochenschr., 109, 293, 1997. 82. Poyart, C., Quesnes, G., and Trieu-Cuot, P., Sequencing the gene encoding manganese dependent superoxide dismutase for rapid species identification of enterococci, J. Clin. Microbiol., 38, 415, 2000. 83. Ke, D.B. et al., Development of a PCR assay for rapid detection of enterococci, J. Clin. Microbiol., 37, 3497, 1999. 84. Wang, R.F. et al., Development of a membrane-array method for the detection of human intestinal bacteria in fecal samples, Mol. Cell. Probes, 16, 341, 2002. 85. Sedgley, C., Buck, G., and Appelbe, O., Prevalence of Enterococcus faecalis at multiple oral sites in endodontic patients using culture and PCR, J. Endodont., 32, 104, 2006. 86. Mahmoudpour, A. et al., Isolation and identification of Enterococcus faecalis from necrotic root canals using multiplex PCR, J. Oral Sci., 49, 221, 2007. 87. Arias, C.A. et al., Rapid identification of Enterococcus hirae and Enterococcus durans by PCR and detection of a homologue of the E. hirae mur 2 gene in E. durans, J. Clin. Microbiol., 44, 1567, 2006. 88. Frahm, E., and Obst, U., Application of the fluorogenic probe technique (TaqMan PCR) to the detection of Enterococcus spp. and Escherichia coli in water samples, J. Microbiol. Methods, 52, 123, 2003.

246 89. Volkmann, H. et al., Detection of clinically relevant antibiotic-resistance genes in municipal wastewater using real-time PCR (TaqMan), J. Microbiol. Methods, 56, 277, 2004. 90. Kempf, V.A J., Trebesius, K., and Autenrieth, I.B. Fluorescent in situ hybridization allows rapid identification of microorganisms in blood cultures, J. Clin. Microbiol., 38, 830, 2000. 91. Waar, K. et al., Fluorescent in situ hybridization with specific DNA probes offers adequate detection of Enterococcus faecalis and Enterococcus faecium in clinical samples, J. Med. Microbiol., 54, 937, 2005. 92. Wellinghausen, N. et al., Rapid identification of clinically relevant Enterococcus species by fluorescence in situ hybridization, J. Clin. Microbiol., 45, 3424, 2007. 93. Betzl, D., Ludwig, W., and Schleifer, K.H., Identification of lactococci and enterococci by colony hybridization with 23S ribosomal RNA targeted oligonucleotide probes, Appl. Environ. Microbiol., 56, 2927, 1990. 94. Lehner, A. et al., Oligonucleotide microarray for identification of Enterococcus species, FEMS Microbiol. Lett., 246, 133, 2005. 95. Frahm, E. et al., Application of 23S rDNA targeted oligonucleotide probes specific for enterococci to water hygiene control, Syst. Appl. Microbiol., 21, 450, 1998. 96. Behr, T. et al., A nested array of rRNA targeted probes for the detection and identification of enterococci by reverse hybridization, Syst. Appl. Microbiol., 23, 563, 2000. 97. Chizhikov, V. et al., Microarray analysis of microbial virulence factors, Appl. Environ. Microbiol., 67, 3258, 2001. 98. Sergeev, N. et al., Multipathogen oligonucleotide microarray for environmental and biodefense applications, Biosens. Bioelectron., 20, 684, 2004. 99. 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. 100. Amann, R.I., Krumholz, L., and Stahl, D.A. Fluorescent oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology, J. Bacteriol., 172, 762, 1990. 101. Jin, L.Q. et al., Detection and identification of intestinal pathogenic bacteria by hybridization to oligonucleotide microarrays, World J. Gastroent., 11, 7615, 2005. 102. Santo Domingo, J.W., Siefring, S.C., and Haugland, R.A., Real time PCR method to detect Enterococcus faecalis in water, Biotechnol. Lett., 25, 261, 2003. 103. Wang, R.F. et al., DNA microarray analysis of predominant human intestinal bacteria in fecal samples, Mol. Cell. Probes, 18, 223, 2004. 104. Tung, S.K. et al., Array based identification of species of the genera Abiotrophia, Enterococcus, Granulicatella, and Streptococcus, J. Clin. Microbiol., 44, 4414, 2006. 105. Manero, A., and Blanch, A.R., Identification of Enterococcus spp. based on specific hybridisation with 16S rDNA probes, J. Microbiol. Methods, 50, 115, 2002. 106. Naser, S.M. et al., Application of multilocus sequence analysis (MLSA) for rapid identification of Enterococcus species based on rpoA and pheS genes, Microbiology, 151, 2141, 2005. 107. Naser, S. et al., Phylogeny and identification of enterococci by atpA gene sequence analysis, J. Clin. Microbiol., 43, 2224, 2005. 108. Jackson, C.R., Fedorka-Cray, P.J., and Barrett, J.B., Use of a genus- and species-specific multiplex PCR for identification of enterococci, J. Clin. Microbiol., 42, 3558, 2004.

Molecular Detection of Human Bacterial Pathogens 109. Goh, S.H. et al., Identification of Staphylococcus species and subspecies by the chaperonin 60 gene identification method and reverse checkerboard hybridization, J. Clin. Microbiol., 35, 3116, 1997. 110. Teng, L.J. et al., Determination of Enterococcus faecalis groESL full length sequence and application for species identification, J. Clin. Microbiol., 39, 3326, 2001. 111. Van de Klundert, J.A.M., and Vliegenthart, J.S., PCR detection of genes coding for aminoglycoside-modifying enzymes. In Persing, D.H. et al. (Editors), Diagnostic Molecular Microbiology, pp. 547–552, ASM Press, Washington, D.C., 1993. 112. Kariyama, R. et al., Simple and reliable multiplex PCR assay for surveillance isolates of vancomycin-resistant enterococci, J. Clin. Microbiol., 38, 3092, 2000. 113. McClellan, D.L., Griffen, A.L., and Leys, E.J. Age and prevalence of Porphyromonas gingivalis in children, J. Clin. Microbiol., 34, 2017, 1996. 114. 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. 115. Nadkarni, M.A. et al., Determination of bacterial load by realtime PCR using a broad-range (universal) probe and primers set, Microbiology, 148, 257, 2002. 116. 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. 117. Kumar, P.S. et al., New bacterial species associated with chronic periodontitis, J. Dent. Res., 82, 338, 2003. 118. 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. 119. Deasy, B.M. et al., A rapid PCR based method to distinguish between Lactococcus and Enterococcus, Syst. Appl. Microbiol., 23, 510, 2000. 120. Matsuda, K. et al., Sensitive quantitative detection of commensal bacteria by rRNA-Targeted reverse TranscriptionPCR, Appl. Environ. Microbiol., 73, 6695, 2007. 121. Matsuda, K. et al., Establishment of an analytical system for the human fecal microbiota, based on reverse transcriptionquantitative PCR targeting of multicopy rRNA molecules, Appl. Environ. Microbiol., 75, 1961, 2009. 122. Barry, T., The 16S723S ribossomal spacer region for DNA probes to identify eubacteria, PCR Methods Appl., 1, 51, 1991. 123. 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. 124. Molander, A. et al., A protocol for polymerase chain reaction detection of Enterococcus faecalis and Enterococcus faecium from the root canal, Int. Endod. J., 35, 1, 2002. 125. 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. 126. 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.

Enterococcus 127. Siqueira, J.F., and Rocas, I.N., Polymerase chain reaction based analysis of microrganisms associated with failed endodontic treatment, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endodontol., 97, 85, 2004. 128. Gomes, B.P.F.A. et al., Enterococcus faecalis in dental root canals detected by culture and by polymerase chain reaction analysis, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endodontol., 102, 247, 2006. 129. Depardieu, F., Perichon, B., and Courvalin, P., Detection of the van alphabet and identification of enterococci and staphylococci at the species level by multiplex PCR, J. Clin. Microbiol., 42, 5857, 2004. 130. Liu, D. et al., PCR amplification of a species-specific putative transcriptional regulator gene reveals the identity of Enterococcus faecalis, Res. Microbiol., 156, 944, 2005. 131. Fines, M. et al., VanE, a new type of acquired glycopeptide resistance in Enterococcus faecalis BM4405, Antimicrob. Agents Chemother., 43, 2161, 1999. 132. McKessar, S.J. et al., Genetic characterization of vanG, a novel vancomycin resistance locus of Enterococcus faecalis, Antimicrob. Agents Chemother., 44, 3224, 2000. 133. Clark, N.C. et al., Characterization of glycopeptide-resistant enterococci from United States hospitals, Antimicrob. Agents Chemother., 37, 2311, 1993. 134. 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. 135. Gold, H.S. et al., A Gene conferring resistance to vancomycin but not teicoplanin in isolates of Enterococcus-faecalis and Enterococcus-faecium demonstrates homology with VanB, VanA, and VanC genes of enterococci, Antimicrob. Agents Chemoter., 37, 1904, 1993. 136. Perichon, B., Reynolds, P., and Courvalin, P., VanD type glycopeptide-resistant Enterococcus faecium BM4339, Antimicrob. Agents Chemother., 141, 2016, 1997. 137. Boyd, D.A. et al., Enterococcus faecium N03–0072 carries a new VanD-type vancomycin resistance determinant: Characterization of the VanD5 operon, J. Antimicrob. Chemother., 54, 680, 2004. 138. Cheng, S.Q. et al., A PCR assay for identification of Enterococcus faecium, J. Clin. Microbiol., 35, 1248, 1997. 139. Satake, S. et al., Detection of vancomycin-resistant enterococci in fecal samples by PCR, J. Clin. Microbiol., 35, 2325, 1997. 140. Semedo-Lemsaddek, T. et al., Enterococcus. In Liu, D. (Editor), Molecular Detection of Foodborne Pathogens, p. 157, CRC Press, Taylor & Fancis Group, New York, 2009. 141. Grunberg-Manago, M., Regulation of the exoression of aminoacyl tRNA synthetases and translation factors. In Neidhardt, F. et al. (Editors), Escherichia coli and Salmonella: Cellular and Molecular Biology, 2nd ed., pp. 1386–1391, ASM Press, Washington, D.C., 1996. 142. Knijff, E. et al., Rapid identification of Enterococcus durans and Enterococcus hirae by PCR with primers targeted to the ddl genes, J. Microbiol. Methods, 47, 35, 2001. 143. Robbi, C. et al., The gene encoding for penicillin-binding protein 5 of Enterococcus faecalis is useful for development of a species-specific DNA probe, Microb. Drug Resist., 2, 215, 1996. 144. Muller, V., and Gruber, G., ATP synthases: structure, function and evolution of unique energy converters, Cell. Mol. Life Sci., 60, 474, 2003.

247 145. Ellis, R.J., Champeronins: introductory perspective. In Ellis, R.J. (Editor), The Chaperonins, pp. 2–25, Academic Press, Inc., San Diego, CA, 1996. 146. Li, S., Zhang, Z., and Mi, Z.H., Vancomycin-resistant enterococci in a Chinese hospital, Curr. Microbiol., 55, 125, 2007. 147. Arthur, M., Reynolds, P., and Courvalin, P., Glycopeptide resistance in enterococci, Trends Microbiol., 4, 401, 1996. 148. Mendéz-Alvarez, S., Pérez-Hernández, X., and ClaverieMartin, F., Glycopeptide resistance in enterococci, Int. Microbiol., 3, 71, 2000. 149. Evers, S., Sahm, D.F., and Courvalin, P., The vanB gene of vancomycin-resistant Enterococcus faecalis V583 is structurally related to genes encoding D-Ala D-Ala ligases and glycopeptide-resistance proteins VanA and VanC, Gene, 124, 143, 1993. 150. Ostrowsky, B.E. et al., A cluster of VanD vancomycin-resistant Enterococcus faecium: Molecular characterization and clinical epidemiology, J. Infect. Dis., 180, 1177, 1999. 151. Linscott, A.J., Collection, transport, and manipulation of clinical specimens and initial laboratory concerns. In Garcia, L.S., and Isenberg, H.D. (Editors in Chief), Clinical Microbiology Procedures Handbook, Volume 1, 2nd ed., pp. 1–36, ASM, Washington D.C., 2007. 152. Thompson, R.B. Jr., Specimen collection, transport, and processing: Bacteriology. In Murray, P.R. et al. (Editors), Manual of Clinical Microbiology, Volume 1, 9th ed., pp. 291–333, ASM, Washington, D.C., 2007. 153. York, M.K., Aerobic bacteriology. In Garcia, L.S., and Isenberg, H.D. (Editors in Chief), Clinical Microbiology Procedures Handbook, Volume 1, 2nd ed., p. 37, ASM, Washington D.C., 2007. 154. Reuter, G., Culture media for enterococci and group D-streptococci. In Corry J.E.L. et al. (Editors), Handbook of Culture Media for Food Microbiology, Volume 34, pp. 51–62, Elsevier Sciences B.V., Amsterdam, The Netherlands, 1995. 155. Reuter, G., and Klein, G., Culture media for enterococci and group D-streptococci. In Corry J.E.L. et al. (Editors), Handbook of Culture Media for Food Microbiology, Volume 37, pp. 111–126, Elsevier Sciences B.V., Amsterdam, The Netherlands, 2003. 156. Atlas, R.M., Handbook of Microbial Media for the Examination of Food, CRC Press, USA, 1995. 157. Solway, S. et al., Isolation of streptogramin resistant Enterococcus faecium from human and non human sources in a rural community, J. Antimicrob. Chemoth., 52, 707, 2003. 158. Swenson, J.M. et al., Development of a standardized screening method for detection of vancomycin-resistant enterococci, J. Clin. Microbiol., 32, 1700, 1994. 159. Asir, K. et al., Evaluation of chromogenic media for the isolation of vancomycin-resistant enterococci from stool samples, Lett. Appl. Microbiol., 48, 230, 2009. 160. Kuch, A. et al., New selective and differential chromogenic agar medium, chromID VRE, for screening vancomycin-resistant Enterococcus species, J. Microbiol. Methods., 77, 124, 2009. 161. Ledeboer, N.A., Tibbetts, R.J., and Dunne, W.A. A new chromogenic agar medium, chromID VRE, to screen for vancomycin-resistant Enterococcus faecium and Enterococcus faecalis, Diagn. Microbiol. Infect. Dis., 59, 477, 2007. 162. Cuzon, G. et al., A novel chromogenic medium for vancomycin resistant Enterococcus faecium and Enterococcus faecalis detection, Int. J. Antimicrob. Agents, 29, S22, 2007. 163. Hughes, J.G. et al., Culture with BACTEC Peds Plus/F bottle compared with conventional methods for detection of bacteria in synovial fluid, J. Clin. Microbiol., 39, 4468, 2001.

248 164. Qian, Q. et al., Direct identification of bacteria from positive blood cultures by amplification and sequencing of the 16S rRNA gene: Evaluation of BACTEC 9240 instrument true positive and false positive results, J. Clin. Microbiol., 39, 3578, 2001. 165. Hamilton-Miller, J.M.T., and Shah, S., Identification of clinically isolated vancomycin-resistant enterococci: Comparison of API and BBL Crystal systems, J. Med. Microbiol., 48, 695, 1999. 166. Peters, R.P.H. et al., Rapid identification of pathogens in blood cultures with a modified fluorescence in situ hybridization assay, J. Clin. Microbiol., 44, 4186, 2006. 167. Miyoung, K. et al., Comparison of the MicroScan, VITEK 2, and Crystal GP with 16S rRNA sequencing and MicroSeq 500 v2.0 analysis for coagulase-negative staphylococci, BMC Microbiol., 8, 233, 2008. 168. Pendle, S. et al., Difficulties in detection and identification of Enterococcus faecium with low-level inducible resistance to vancomycin, during a hospital outbreak, Clin. Microbiol. Infec., 14, 853, 2008. 169. Vankerckhoven, V. et al., Development of a multiplex PCR for the detection of asa1, gelE, cylA, esp, and hyl genes in enterococci and survey for virulence determinants among European hospital isolates of Enterococcus faecium, J. Clin. Microbiol., 42, 4473, 2004. 170. Chen, A.Y., and Zervos, M.J., Enterococcus: Antimicrobial resistance in enterococci epidemology, treatment, and control. In Mayers, D.L. (Editor), Antimicrobial Drug Resistance, pp. 715–734, Humana Press, 2009.

Molecular Detection of Human Bacterial Pathogens 171. Centers for Disease Control and Prevention. Recommendations for Preventing the Spread of Vancomycin Resistance Recommendations of the Hospital Infection Control Practices Advisory Committee (HICPAC). MMWR, 44, 1, 1995. 172. Owens, R.C. Jr., and Ambrose, P.G., Antimicrobial stewardship and the role of pharmacokinetics-pharmacodynamics in the modern antibiotic era, Diagn. Microbiol. Infect. Dis., 57, 77S, 2007. 173. Owens, R.C. Jr., Antimicrobial stewardship: concepts and strategies in the 21st century, Diagn. Microbiol. Infect. Dis., 61, 110, 2008. 174. Patel, D., Lawson, W., and Guglielmo, B.J., Antimicrobial stewardship programs: Interventions and associated outcomes, Expert Rev. Anti. Infect. Ther., 6, 209, 2008. 175. Dellit, T. et al., Infectious Diseases Society of America and the Society for Healthcare Epidemiology of America guidelines for developing an institutional program to enhance antimicrobial stewardship, Clin. Infect. Dis., 44, 159, 2007. 176. Drew, R.H., Antimicrobial stewardship programs: How to start and steer a successful program, J. Manag.Care Pharm., 15, S18, 2009. 177. Brown, M., and McClure, P., Microbiological risk assessment for emerging pathogens. In Motarjemi, Y., and Adams, M. (Editors), Emerging Foodborne Pathogens, pp. 130–152, Woodhead Publishing Limited and CRC Press LLC, England, 2006. 178. Skovgaard, N., New trends in emerging pathogens, Int. J. Food Microbiol., 120, 217, 2007.

23 Granulicatella Sheng Kai Tung and Tsung Chain Chang CONTENTS 23.1 Introduction...................................................................................................................................................................... 249 23.1.1 Classification, Morphology, and Biology............................................................................................................. 249 23.1.2 Clinical Features................................................................................................................................................... 249 23.1.3 Diagnosis.............................................................................................................................................................. 250 23.1.3.1 Conventional Techniques....................................................................................................................... 250 23.1.3.2 Molecular Techniques............................................................................................................................ 251 23.2 Methods............................................................................................................................................................................ 251 23.2.1 Sample Preparation............................................................................................................................................... 251 23.2.2 Detection Procedures............................................................................................................................................ 251 23.2.2.1 16S rRNA Gene Sequencing.................................................................................................................. 251 23.2.2.2 Sequencing of the Ribosomal 16S-23S Intergenic Spacer (ITS) Region............................................... 252 23.2.2.3 Partial rpoB Gene Sequencing.............................................................................................................. 252 23.2.2.4 Array-Based Method.............................................................................................................................. 252 23.3 Conclusion and Future Perspectives................................................................................................................................. 253 References.................................................................................................................................................................................. 254

23.1  INTRODUCTION 23.1.1  Classification, Morphology, and Biology Bacteria in the genus Granulicatella are gram-positive, catalase-negative, facultative anaerobic cocci. Species in the genus can cause opportunistic infections in immunocompromised patients and are infrequently isolated from clinical laboratories.1 The genera Abiotrophia and Granulicatella have been proposed to accommodate microorganisms previously known as nutritionally variant streptococci or satelliting streptococci.1–3 By chromosomal DNA–DNA hybridizations, Bouvet et al.4 demonstrated that nutritionally variant streptococci were really members of two novel streptococcal species, Streptoccoccus defectivus and Streptoccoccus adjacens. Based on the 16S rRNA gene sequence, Kawamura et al.3 transferred Streptoccoccus defectivus and Streptoccoccus adjacens, respectfully, to two new species, Abiotrophia defectiva and Abiotrophia adiacens. A third species, Abiotrophia elegans, a possible pathogen in patients with endocarditis, was described in 1998 by Roggenkamp et al.5 In 2000, Collins and Lawson2 proposed a new genus, Granulicatella, with Granulicatella adiacens and Granulicatella elegans encompassing strains formerly recognized as Abiotrophia adiacens and Abiotrophia elegans, respectfully. Abiotrophia defectiva remains as the sole species in Abiotrophia.2 A third species in Granulicatella is Granulicatella balaenopterae; however, there was no report of human infection caused by this microorganism.

Bacteria in the genus Granulicatella are nonmotile cocci that occur as single cells, in pairs, or in short chains. Cell morphology depends on culture conditions. Elongated or swollen cells may be observed, especially when grown under less optimal nutritional conditions.6 The G + C content of Granulicatella DNA is approximately 36–37.5 mol%. Members of Granulicatella are normal flora of the oral cavity or upper respiratory tract1 and are non-betahemolytic on blood-based solid media. Granulicatella exhibit satellitism around the colonies of other bacteria, such as Staphylococcus epidermidis and other streptococci.7 Granulicatella species usually have low virulence and are normally pathogenic in immunocompromised hosts.1 Predisposing factors that lead to Granulicatella infections include previously damaged tissues (e.g., heart valves), invasive procedures, prolonged hospitalization, and the presence of foreign bodies. Granulicatella can be isolated from a variety of clinical samples, including blood, urine, cerebrospinal fluid, wound tissues, and others specimens.1 Infections caused by Granulicatella might be due to some microorganisms entering the bloodstream and disseminating to other tissues.8,9

23.1.2  Clinical Features Granulicatella species have been reported as etiologic agents of bacteremia, infectious endocarditis, brain abscess, postpartum or postabortal sepsis, pancreatic abscess, wound 249

250

infection, vertebral osteomyelitis, conjunctivitis, cirrhosis, endophthalmitis, infectious crystalline keratopathy, septic arthritis, otitis media, central nervous system infections, breast implant–associated infection, and peritoneal dialysis– related peritonitis.5,7,10–17 Using a molecular technique, the proportion of Granulicatella elegans, Granulicatella adiacens, and Abiotrophia defectiva was found to be 1:11:1 in the human mouth.18 Exopolysaccharides that can adhere to human tissues are produced by Granulicatella.7 All members of Granulicatella are susceptible to vancomycin. However, there are still no standardized methods and interpretation criteria for antibiotic susceptibility testing results in the documents of CLSI (Clinical and Laboratory Standards Institute). Most investigators have employed streptococcal (other than Streptococcus pneumoniae) interpretive criteria for determining the results of susceptibility.1 Since Granulicatella species are fastidious, Mueller-Hinton medium supplemented with blood and pyridoxal hydrochloride (0.001%) is used for susceptibility testing, and a CO2-enriched atmosphere1 is required for incubation. Tuohy et al.19 examined a collection of 21 Granulicatella adiacens isolates for their susceptibilities to a variety of antibiotics. They found that all strains were susceptible to clindamycin, rifampin, levofloxacin, ofloxacin, and quinupristin-dalfopristin. The percentage of isolates that were susceptible to other agents was as follows: penicillin, 55%; amoxicillin, 81%; ceftriazone, 63%; and meropenem, 96%. However, Zheng et al.20 observed high rates of beta-lactam and macrolide resistance in a collection of pediatric Granulicatella isolates, and hence they proposed that antimicrobial susceptibility testing should be systematically done to achieve appropriate antimicrobial therapy. In severely ill patients or those with a suboptimal response to initial therapy with beta-lactam antibiotics, treatment with vancomycin should be considered.21 The combination therapy with vancomycin and an aminoglycoside should be initiated empirically in patients with central nervous system infections.13

23.1.3  Diagnosis 23.1.3.1  Conventional Techniques Phenotypic characteristics that can be used as a guide to identify Granulicatella and Abiotrophia species are listed in Table 23.1. Members of Granulicatella are likely to be isolated on rich and nonselective media. They can grow on chocolate agar, in thioglycolate broth, and on brucella agar with 5% horse blood, but not on Typticase soy agar with 5% sheep blood.1 Cell growth requires thiol compounds such as l-cysteine or the active form of vitamin B6, pyridoxal phosphate, in media. Growth does not occur at 10°C or 45°C. Granulicatella in clinical samples may be overlooked when media without nutritional supplements are used. Gram stain morphology, although prone to be subjective in interpretation, is a major decision point in the identification protocols. Gram stain morphology of Granulicatella spp. resembles that of streptococci, with cells being cocci or coccobacilli in pairs and chains. Cells grown in broth, such as thioglycollate, should be used for determination of cellular morphology.

Molecular Detection of Human Bacterial Pathogens

Besides the characteristics listed in Table 23.1, additional tests are recommended to make a final identification, especially for isolates recovered from important clinical specimens.1 Additional tests useful for Granulicatella spp. differentiation can be found in several studies.4,6,22 Commercial identification kits, such as API 20 Strep, rapid ID32 STREP, and the VITEK 2 system (all from BioMérieux, Marcy l’Etoile, France), could be used for identification of Granulicatella spp. However, Granulicatella elegans cannot be identified by both API 20 Strep and rapid ID32 STREP kits since the species is not included in the databases of the two kits. Granulicatella adiacens and Granulicatella elegans are phenotypically similar; several tests (hydrolysis of arginine and hippurate, and production of β-glucuronidase) can be used to differentiate the two species (Table 23.1). Most clinical Granulicatella adiacens isolates could be identified by the API 20 STREP system with > 80%–90% confidence, whereas both the Vitek System (GPI) and ID32 STREP identified only a small portion of Granulicatella adiacens isolates.23 It should be noted that the growth of Granulicatella elegans is only supported by l-cysteine hydrochloride and not by pyridoxal phosphate when the organism is grown in ToddHewitt or casein-soy peptone broth.5 The media of blood culture systems supplemented only with pyridoxal phosphate TABLE 23.1 Phenotypic Characteristics of Abiotrophia defectiva and Granulicatella Speciesa Characteristics A. defectiva G. adiacens G. balaenopterae G. elegans Production of acid from Lactose Maltose Pullulan Sucrose Tagatose Trehalose

+ + + + − +

− + − + + −

− + + − − +

− − − + − −

Hydrolysis of Arginine Hippurate

− −

− −

+ −

+ +

Production of a-Galactosidase

+

−

−

−

β-Galactosidase

+

−

−

−

β-Glucuronidase

−

+

−

−

N-Acetyl-βglucosaminidase Pyrrolidonyl arylamidase Leucine aminopeptidase Satelliting behavior

−

−

+

−

+

+

ND

+

+

+

ND

+

+

+

ND

+

Symbols: −, negative reactions ≤15%; +, positive reactions ≥85%; ND, not determined. a  Data compiled from References 1, 2, 5, 6, 44, and 45.

251

Granulicatella

may fail to support the growth of Granulicatella elegans, and thus, these systems might not be able to detect this microorganism as a possible pathogen. 23.1.3.2  Molecular Techniques Molecular methods developed for identification of members of Granulicatella are limited. Sequence analysis of the 16S rRNA gene has been widely used for bacterial identification.24,25 The technique is becoming popular for identifying biochemically unidentified bacteria or for providing reference identifications for unusual strains.24 Sequencing of the 16S rRNA gene can be successfully used to identify Granulicatella species. Multiple sequences of the 16S rRNA genes for Granulicatella adiacens and Granulicatella elegans are available in the nucleotide database of National Center for Biotechnology Information, USA. The 16S rRNA gene sequences of Granulicatella adiacens and Abiotrophia defectiva formed a distinct cluster; this cluster is not closely related to other viridans streptococcal species and Abiotrophia.3,5 Woo et al.23 found that 16S rRNA gene sequencing is the technique of choice for identifying Granulicatella adiacens and proposed that early surgical intervention should be considered when endocarditis caused by this microorganism is diagnosed. The ribosomal 16S-23S intergenic spacer (ITS) region has been suggested as a good candidate for bacterial identification and strain typing.26–29 Sequences of the ITS region in bacteria have low intraspecies variation and high interspecies divergence.26,27,29 The feasibility of using ITS sequence to identify Granulicatella adiacens and Granulicatella elegans was established by Tung et al.30 It should be noted that in the genome of Granulicatella spp. possesses multiple ITS fragments having different lengths and sequences, therefore the PCR products amplified from the ITS region of an isolate cannot be directly sequenced. Before sequencing, the amplicons should be eluted from agarose gel or cloned into a plasmid and reamplified. Once the ITS sequence is determined, species identification can be done by searching databases using the BLAST (http://www.ncbi.nlm.nih.gov/ blast/) sequence analysis tool. The rpoB gene, encoding the highly conserved subunit of the bacterial RNA polymerase, has been demonstrated to be a suitable target for identification of a wide range of bacteria.31 The gene was shown to be more discriminative than the 16S rRNA gene for identifying enteric bacteria.31 Partial rpoB gene sequence (740 bp) had been applied to identify streptococci and Granulicatella adiacens.32 In recent years, DNA array technology has been successfully used to identify microorganisms that are difficult to be determined to the species level; these bacteria include viridans streptococci,33 nonfermenting gram-negative bac­ teria,34 Legionella,35 Mycobacterium36,37 and foodborne bacterial pathogens.38 The method consists of PCR amplification of a specific gene region, followed by hybridization of the PCR product labeled with a fluorescent dye (or with a molecular technique that could be recognized by antibody) to a panel of oligonucleotide probes immobilized on a solid

support such as glass slide or nylon membrane. The test sensitivity and specificity of the arrays are largely determined by the sequences of the oligonucleotide probes. The whole procedure of array hybridization takes between 8 and 24 h, starting from isolated colonies or clinical specimens. The hybridization patterns could be read by a fluorescence scanner if a fluorescent label is used or by the naked eye if colorimetric reaction is used. By using the oligonucleotide array platform, five oligonucleotide probes designed from the ITS regions were successfully used to identify Granulicatella adiacens, Granulicatella elegans, and Granulicatella balaenopterae.39 The array method used a protocol encompassing DNA extraction, PCR amplification of the ITS regions, and hybridization of the PCR products to speciesspecific probes on the array.

23.2  METHODS 23.2.1  Sample Preparation The boiling method described by Millar et al. could be used to extract DNA from the bacterial colonies.40 Briefly, one to several colonies from pure cultures are suspended in 50 µL of sterilized water and heated at 100°C for 15 min in a heating block. After centrifugation in a microcentrifuge (6000 × g for 3 min), the supernatant contains bacterial DNA. Alternatively, a variety of DNA extraction kits may be also used.

23.2.2  Detection Procedures 23.2.2.1  16S rRNA Gene Sequencing Principle.  Relman25 utilized primer pair 8FPL (5′-GTTT GATCCTGGCTCAG-3′) and 1492RPL (5′-GGTTACCT TGTTACGACTT-3′) for PCR amplification of the nearly complete length of the 16S rRNA gene. Sequencing analysis of the resulting fragment reveals the identity of the bacteria under investigation. Procedure 1. Prepare PCR mixture (50 µL) consisting of 10 mM Tris-HCl pH 8.3, 50 mM KCl, 1.5 mM MgCl2, 1 mM dNTPs (0.25 mM each), 0.1 µM (each) primer, 1 U of Taq DNA polymerase, and 5 µL (1–5 ng) of template DNA. 2. Conduct PCR amplification with 1 cycle of 94°C for 5 min; 5 cycles of 94°C for 1 min, 52°C for 1 min, and 72°C for 1.5 min; 25 cycles of 94°C for 1 min, 55°C for 1 min, and 72°C for 1.5 min; and a final 72°C for 5 min. 3. Purify and sequence PCR products in both directions by using the above two primers and an additional primer, 1055r (5′-CACGAGCTGACGACAG CCAT-3′).25,27 4. Compare the determined sequences to known sequences of the 16S rRNA genes in the databases of the National Center for Biotechnology Information using the BLASTN algorithm.

252

Note: Species identification is determined from the bestscoring reference sequence of the BLAST output that has a sequence similarity of ≥99% with the query sequence. All positions showing differences from the best-scoring reference sequence could be visually inspected in the electropherogram, and the sequence is corrected if needed, that is, when obvious sequencing software errors occur, or when undetermined nucleotides in the sequence can be determined by visual inspection of the electropherogram. Afterwards, a second search is done with the BLAST algorithm. Ideally, 1300–1500 bp are sequenced with 99.5% sequence similarity, with sequence match to type strain or reference strain of species that has undergone DNA-relatedness studies. 23.2.2.2  S equencing of the Ribosomal 16S-23S Intergenic Spacer (ITS) Region Principle.  Gürtler and Stanisich28 designed the bacteriaspecific universal primer pair 13BF (5′-GTGAATACGTTC CCGGGCCT-3′) and 6R (5′-GGGTTYCCCCRTTCR GAAAT-3′) (where Y is C or T and R is A or G) for amplification of a DNA fragment that encompassed ITS, a portion of the 3′ end of the 16S rRNA gene, and a portion of the 5′ end of the 23S rRNA gene.27,30 The 5′ end of primer 13BF is located at position 1371 of the 16S rRNA gene, and the 5′ end of primer 6R is located at position 108 downstream of the 5′ end of the 23S rRNA gene (Escherichia coli numbering). The identity of the bacteria concerned is determined following nucleotide sequencing analysis of the ITS fragment. Procedure 1. Prepare PCR mixture (50 µL) consisting of 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.8 mM deoxyribonucleoside triphosphates (0.2 mM each), 0.1 µM primer (each), 1 U of Taq DNA polymerase, and 5 µL (1–5 ng) of template DNA. 2. Perform PCR amplification with 1 cycle of 94°C for 5 min; 5 cycles 94°C for 1 min; 55°C for 1 min, and 72°C for 1 min; 30 cycles of 94°C for 1 min, 60°C for 1 min, and 72°C for 1 min; and a final 72°C for 3 min. 3. Purify and sequence PCR products in both directions by using the above two primers. Note: Species of Granulicatella produce multiple ITS amplicons in PCR with different length and sequence.27 Only the shortest ITS fragment of a strain, which is usually the dominant band, is eluted from the agarose gel and sequenced. In this way, the ITS of Granulicatella elegans can be sequenced. However, the amplicons of Granulicatella adiacens eluted from gels cannot be directly sequenced due to the presence of different amplicons with similar size in the eluted DNA. For this reason, the eluted DNA from gel can be cloned with a Topo TA cloning kit (Invitrogen, Carlsbad, California, USA). The ITS regions of positive clones are reamplified and sequenced. The determined sequences are compared to

Molecular Detection of Human Bacterial Pathogens

known sequences in the databases of the National Center for Biotechnology Information using the BLASTN algorithm. Species identity is determined from the best-scoring reference sequence of the BLAST output and whether the best-scoring reference sequence in the databases has a sequence identity of ≥98% with the query sequence.27 23.2.2.3  Partial rpoB Gene Sequencing Principle.  Drancourt et al.32 employed primers Strepto F (5′-AARYTIGGMCCTGAAGAAAT-3′) and Strepto R (5′-T GIARTTTRTCATCAACCATGTG-3′) (where M is A or C, R is A or G, and W is A or T) to amplify a portion (740 bp) of the rpoB gene from Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella. Procedure 1. Prepare PCR mixture (50 µL) consisting of 2.5 U of Taq polymerase, 1× Taq buffer, 1.8 mM MgCl2 (Gibco-BRL/Life Technologies, Cergy Pontoise, France), 0.2 mM concentrations of dATP, dTTP, dGTP, and dCTP, and 0.2 μM concentrations of each primer, and 5 µL (1–5 ng) of template DNA. 2. Subject the PCR mixture to one cycle of 95°C for 2 min; 35 cycles of 94°C for 30 s, 52°C for 30 s, and 72°C for 60 s; and a final 72°C for 5 min. 3. Purify the amplified product by using a QIAquick spin PCR purification kit (Qiagen, Hilden, Germany) and determine the nucleotide sequence of the amplicon. Note: This primer pair Strepto F and Strepto R is shown to be almost specific for streptococcal species, since no amplification products were obtained from 58 other bacterial isolates belonging to nonstreptococcal species, including other grampositive cocci.32 23.2.2.4  Array-Based Method Principle.  Tung et al.39 reported an array-based method for identification of Abiotrophia (1 species), Enterococcus (18 species), Granulicatella (3 species), and Streptococcus (31 species and 6 subspecies). The method consists of PCR amplification of the ribosomal DNA intergenic spacer (ITS) regions, and hybridization of the digoxigenin-labeled PCR products to a panel of 88 oligonucleotide probes (16- to 30-mers) immobilized on a nylon membrane. The probes are designed either from the ITS regions or from the 3′ ends of the 16S rRNA genes, and belong to three categories: species specific, group specific, and supplemental probes. Procedure 1. Amplify the ITS region for hybridization with primer pair 8FPL (5′-GTTTGATCCTGGCTCAG-3′) and 1492RPL (5′-GGTTACCTTGTTACGACTT-3′) as shown in Section 23.2.2.2, except that the reverse primer 6R (5′dig-GGGTTYCCCCRTTCRGAAAT3′) is labeled with a digoxigenin molecule at its 5′ end.39 Labeling of both primers with digoxigenin is encouraged because this can increase the hybri­dization signal.

253

Granulicatella

















2. Fabricate oligonucleotide array: (a) Customer synthesize Granulicatella adiacens– specific probe (5′-TATCACAACAAATAACC AATTAA-3′), Granulicatella elegans probes (5′- GAG GT TA AC TC TCA AC TCGACC T TTGAAAA-3′, 5′-TTTAACAAGAAGCAACG CGACCATA-3′) and Granulicatella balaenopterae-specific probes (5′-ATAAC GGA ACCTACCAAGTTCACTTC-3′, 5′-TGAGAGA T TA AT T C T C T C TAGAC T T T GAT C -3 ′, 5′- G CA A ACG CGA AT CATAT TGAGAC TTAA-3′)30; (b) Add multiple thymine bases (5–15) to the 3′ ends of the above probes to increase the binding of these probes to the nylon membrane that is a solid support for immobilizing probes41; (c) Design a positive control probe (5′-GTCGT AACAAGGTAGCCGTA-3′) from a conserved region at the 3′ end of the 16S rRNA gene (GenBank AB023575) to verify all the procedures including PCR, hybridization, antibody reaction, and colorimetric reaction; (d) Dilute the oligonucleotide probes 1:1 (final concentration, 10 µM) with a tracking dye solution (30% [vol/vol] glycerol, 40% [vol/vol] dimethyl sulfoxide, 1 mM EDTA [disodium salt], 0.15% [wt/vol] bromophenol blue, 10 mM Tris-HCl [pH 7.5]). (e) Draw the probe solutions into different wells of a round-bottom microtiter plate and spotted onto a positively charged nylon membrane (Roche, Mannheim, Germany) with an automatic arrayer (SR-A300; Ezspot, Taipei, Taiwan) by using a solid pin (400–1000 µm in diameter); alternatively, spot the probe solutions on the membrane using the solid pin by hand; use dye solution without probe as a negative control; (f) Dry and expose the membrane to short-wave UV light (Stratalinker 1800, California, USA) for 30 s; Remove unbound oligonucleotides by two washes (2 min each) at room temperature in 0.5 × SSC (1 × SSC is 0.15 M NaCl plus 0.015 M sodium citrate)-0.1% sodium dodecyl sulfate (SDS); (g) Air dry and store the arrays at room temperature under desiccation for further use. 3. Hybridize the labeled PCR products to probes: (a) Prehybridize each array for 2 h with 1 mL of hybridization solution (5 × SSC, 1% [wt/vol] blocking reagent (Roche), 0.1% N-laurylsarcosine,  0.02% SDS) in an individual well of a 24-well cell culture plate. (b) Heat the digoxigenin-labeled PCR product amplified from an isolate in a boiling water bath for 5 min and immediately cool on an ice bath. (c) Hybridize at 45°C for 90 min in an incubator with a shaking speed of 60 rpm.











(d) Wash the array three times (5 min each) in 1 mL of washing buffer (2× SSC, 0.1% SDS) and one wash (1 min) in 1 mL of a second washing buffer (0.5× SSC, 0.1% SDS). (e) Block for 1 h with 1 mL of blocking solution supplied in the DIG Nucleic Acid Detection kit (Roche) (dissolve 1% [wt/vol] blocking reagent in maleic acid buffer [0.1 M maleic acid, 0.15 M NaCl, pH 7.5]). (f) Remove the blocking solution, add 0.5 mL of alkaline phosphatase-conjugated sheep antidigoxigenin antibodies (diluted 1:2500 in blocking solution) to each well, and incubate for 1 h. (g) Wash the array three times (15 min each) in 1 mL of washing solution (0.3% [vol/vol] Tween 20 in maleic acid buffer), wash once in 1 mL of detection buffer (0.1 M Tris-HCl, 0.15 M NaCl, pH 9.5) for 5 min. (h) Add 0.5 mL of alkaline phosphatase substrate (a  stock solution of nitroblue tetrazolium chloride and 5-bromo-4-chloro-3-indolylphosphate diluted 1:50 in detection buffer) (Roche) to each well, and incubate the plate at 37°C in dark (without shaking).

Note: Color development is clearly visible between 30 min and 1 h after the start of the reaction; then the hybridized spot can be easily recognized by the naked eye. A strain is identified as one of the Granlicatella species when the probe (or all probes) specified for that species is hybridized.

23.3  C  ONCLUSION AND FUTURE PERSPECTIVES Members of the genus Granulicatella bacteria are grampositive, catalase-negative, and facultatively anaerobic cocci. Currently, the genus contains three species, Granulicatella adiacens, Granulicatella elegans, and Granulicatella balaenopterae, with the first two being human pathogens. Granulicatella spp. normally display low virulence to hosts and are infrequently isolated as opportunistic pathogens.1 These bacteria may enter the bloodstream and disseminate to other tissues and thus cause infections.8,9 A wide spectrum of infections caused by Granulicatella has been reported.7,10,11,13–17,42,43 They resemble other better known streptococcal isolates and consequently may be misidentified as these bacteria. Granulicatella spp. are part of the normal flora of human pharynx, urogenital, and intestinal tracts.1,7 They form tiny colonies on blood agar supplemented with pyridoxal phosphate or l-cysteine and exhibit satellitism around the colonies of Staphylococcus epidermidis and other streptococci.7 Granulicatella may be overlooked because of their poor growth on unsupplemented media and may not be accurately identified by conventional diagnostic procedures or commercial kits. Molecular techniques are good alternatives for identification of doubtful organisms.

254

Molecular methods used to identify Granulicatella species include 16S rRNA gene sequencing, ribosomal 16S-23S intergenic spacer (ITS) sequencing, partial sequencing of the rpoB gene, and oligonucleotide probe hybridization. Sequencing of the 16S rRNA gene seems to be a prior choice to identify Granulicatella spp. since their respective 16S rRNA gene sequences are available in GenBank. Sequencing of the ribosomal 16S-23S intergenic spacer (ITS) is another alternative. However, Granulicatella spp. possess multiple ITS fragments with different length and sequence; therefore, the amplified ITS should be eluted from agarose gel or cloned before sequencing.30 For this reason, ITS sequencing seems to be impractical for identification of Granulicatella. This drawback can be overcome by the hybridization technique, with probes being designed from the ITS region. Five oligonucleotide probes were successfully used to identify the three Granulicatella species.39 The array format originally developed by Tung et al.39 can be easily transformed to the dot hybridization format without sacrifice of the probe efficacy. Finally, partial sequencing (740 bp) of rpoB, encoding the subunit of the bacterial RNA polymerase, could be used to identify Granulicatella adiacens.32 Unfortunately, there is no rpoB sequence of Granulicatella elegans deposited in public databases. Accurate identification of members of Granulicatella may be of great help in elucidating the epidemiology of these bacteria.

REFERENCES









1. Ruoff, K.L., Aerococcus, Abiotrophia, and other aerobic catalase-negative, gram-positive cocci. In Murray, P. R. et  al. (Editors), Manual of Clinical Microbiology, 9th ed., pp. 443–454, American Society for Microbiology, Washington, DC, 2007. 2. Collins, M.D., and Lawson, P.A., The genus Abiotrophia (Kawamura et al.) is not monophyletic: Proposal of Granulica­ tella gen. nov., Granulicatella adiacens comb. nov., Granulica­ tella elegans comb. nov. and Granulicatella balaenopterae comb. nov., Int. J. Syst. Evol. Microbiol., 50, 365, 2000. 3. Kawamura, Y. et al., Transfer of Streptococcus adjacens and Streptococcus defectivus to Abiotrophia gen. nov. as Abiotrophia adiacens comb. nov. and Abiotrophia defectiva comb. nov., respectively, Int. J. Syst. Bacteriol., 45, 798, 1995. 4. Bouvet, A., Grimont, F., and Grimont, P.A.D., Streptococcus defectivus sp. nov. and Streptococcus adjacens sp. nov., nutritionally variant streptococci from human clinical specimens, Int. J. Syst. Bacteriol., 39, 290, 1989. 5. Roggenkamp, A. et al., Abiotrophia elegans sp. nov., a possible pathogen in patients with culture-negative endocarditis, J. Clin. Microbiol., 36, 100, 1998. 6. Christensen, J.J., and Facklam, R.R., Granulicatella and Abiotrophia species from human clinical specimens, J. Clin. Microbiol., 39, 3520, 2001. 7. Ruoff, K.L., Nutritionally variant streptococci, Clin. Microbiol. Rev., 4, 184, 1991. 8. Coykendall, A.L., Classification and identification of the viridans streptococci, Clin. Microbiol. Rev., 2, 315, 1989. 9. Strom, B.L. et al., Dental and cardiac risk factors for infective endocarditis. A population-based, case-control study, Ann. Intern. Med., 129, 761, 1998.

Molecular Detection of Human Bacterial Pathogens 10. Altay, M. et al., A novel agent of peritoneal dialysis-related peritonitis: Granulicatella adiacens, Perit. Dial. Int., 28, 96, 2008. 11. Biermann, C. et al., Isolation of Abiotrophia adiacens from a brain abscess which developed in a patient after neurosurgery, J. Clin. Microbiol., 37, 769, 1999. 12. Bouvet, A., Human endocarditis due to nutritionally variant streptococci: Streptococcus adjacens and Streptococcus defectivus, Eur. Heart. J., 16 Suppl B, 24, 1995. 13. Cerceo, E. et al., Central nervous system infections due to Abiotrophia and Granulicatella species: An emerging challenge? Diagn. Microbiol. Infect. Dis., 48, 161, 2004. 14. Heath, C.H. et al., Vertebral osteomyelitis and discitis associated with Abiotrophia adiacens (nutritionally variant streptococcus) infection, Aust. N. Z. J. Med., 28, 663, 1998. 15. Hepburn, M.J. et al., Septic arthritis caused by Granulicatella adiacens: Diagnosis by inoculation of synovial fluid into blood culture bottles, Rheumatol. Int., 23, 255, 2003. 16. Namdari, H. et al., Abiotrophia species as a cause of endophthalmitis following cataract extraction, J. Clin. Microbiol., 37, 1564, 1999. 17. Ormerod, L.D. et al., Infectious crystalline keratopathy. Role of nutritionally variant streptococci and other bacterial factors, Ophthalmology, 98, 159, 1991. 18. Sato, S. et al., Abiotrophia elegans strains comprise 8% of the nutritionally variant streptococci isolated from the human mouth, J. Clin. Microbiol., 37, 2553, 1999. 19. Tuohy, M.J. et al., Antimicrobial susceptibility of Abiotrophia adiacens and Abiotrophia defectiva, Diagn. Microbiol. Infect. Dis., 38, 189, 2000. 20. Zheng, X. et al., Antimicrobial susceptibilities of invasive pediatric Abiotrophia and Granulicatella isolates, J. Clin. Microbiol., 42, 4323, 2004. 21. Senn, L. et al., Bloodstream and endovascular infections due to Abiotrophia defectiva and Granulicatella species, BMC Infect. Dis., 6, 9, 2006. 22. Beighton, D. et al., Analysis of enzymatic activities for differentiation of two species of nutritionally variant streptococci, Streptococcus defectivus and Streptococcus adjacens, J. Clin. Microbiol., 33, 1584, 1995. 23. Woo, P.C. et al., Granulicatella adiacens and Abiotrophia defectiva bacteraemia characterized by 16S rRNA gene sequencing, J. Med. Microbiol., 52, 137, 2003. 24. Janda, J.M., and Abbott, S.L., 16S rRNA gene sequencing for bacterial identification in the diagnostic laboratory: Pluses, perils, and pitfalls, J. Clin. Microbiol., 45, 2761, 2007. 25. Relman, D.A., Universal bacterial 16S rDNA amplification and sequencing. In Persing, D.H. et al. (Editors), Diagnostic Molecular Microbiology, pp. 489–495, American Society for Microbiology, Washington, DC, 1993. 26. Chang, H.C. et al., Species-level identification of isolates of the Acinetobacter calcoaceticus–Acinetobacter baumannii complex by sequence analysis of the 16S–23S rRNA gene spacer region, J. Clin. Microbiol., 43, 1632, 2005. 27. Chen, C.C. et al., Identification of clinically relevant viridans group streptococci by sequence analysis of the 16S-23S ribosomal DNA spacer region, J. Clin. Microbiol., 42, 2651, 2004. 28. Gürtler, V., and Stanisich, V.A., New approaches to typing and identification of bacteria using the 16S–23S rDNA spacer region, Microbiology, 142, 3, 1996. 29. Hassan, A.A. et al., Inter- and intraspecies variations of the 16S–23S rDNA intergenic spacer region of various streptococcal species, Syst. Appl. Microbiol., 26, 97, 2003.

Granulicatella 30. Tung, S.K. et al., Identification of species of Abiotrophia, Enterococcus, Granulicatella and Streptococcus by sequence analysis of the ribosomal 16S-23S intergenic spacer region, J. Med. Microbiol., 56, 504, 2007. 31. Mollet, C., Drancourt, M., and Raoult, D., rpoB sequence analysis as a novel basis for bacterial identification, Mol. Microbiol., 26, 1005, 1997. 32. Drancourt, M. et al., rpoB gene sequence-based identification of aerobic gram-positive cocci of the genera Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella, J. Clin. Microbiol., 42, 497, 2004. 33. Chen, C.C. et al., Identification of clinically relevant viridans streptococci by an oligonucleotide array, J. Clin. Microbiol., 43, 1515, 2005. 34. Su, S.C. et al., Identification of nonfermenting Gram-negative bacteria of clinical importance by an oligonucleotide array, J. Med. Microbiol., 58, 596, 2009. 35. Su, H.-P. et al., Identification of Legionella species by an oligonucleotide array, J. Clin. Microbiol., 47, 1386, 2009. 36. Fukushima, M. et al., Detection and identification of Mycobacterium species isolates by DNA microarray, J. Clin. Microbiol., 41, 2605, 2003. 37. Park, H. et al., Detection and genotyping of Mycobacterium species from clinical isolates and specimens by oligonucleotide array, J. Clin. Microbiol., 43, 1782, 2005.

255 38. Lin, M.C. et al., Identification of six foodborne pathogens and Psedomonas aeruginosa grown on selective media by an oligonucleotide array, J. Food Prot., 68, 2278, 2005. 39. Tung, S.K. et al., Array-based identification of species of the genera Abiotrophia, Enterococcus, Granulicatella, and Streptococcus, J. Clin. Microbiol., 44, 4414, 2006. 40. Millar, B.C. et al., A simple and sensitive method to extract bacterial, yeast and fungal DNA from blood culture material, J. Microbiol. Methods, 42, 139, 2000. 41. Brown, T.J., and Anthony, R.M., The addition of low numbers of 3′ thymine bases can be used to improve the hybridization signal of oligonucleotides for use within arrays on nylon supports, J. Microbiol. Methods, 42, 203, 2000. 42. del Pozo, J.L. et al., Granulicatella adiacens breast implant-associated infection, Diagn. Microbiol. Infect. Dis., 61, 58, 2008. 43. Gephart, J.F., and Washington, J.A. 2nd, Antimicrobial susceptibilities of nutritionally variant streptococci, J. Infect. Dis., 146, 536, 1982. 44. Kanamoto, T. et al., Genetic heterogeneities and phenotypic characteristics of strains of the genus Abiotrophia and proposal of Abiotrophia para-adiacens sp. nov., J. Clin. Microbiol., 38, 492, 2000. 45. Ohara-Nemoto, Y. et al., Infective endocarditis caused by Granulicatella elegans originating in the oral cavity, J. Clin. Microbiol., 43, 1405, 2005.

24 Lactobacillus Ester Sánchez and Yolanda Sanz CONTENTS 24.1 Introduction...................................................................................................................................................................... 257 24.1.1 Taxonomy............................................................................................................................................................. 257 24.1.2 Ecology................................................................................................................................................................. 258 24.1.3 Phenotypic Characterization................................................................................................................................ 258 24.1.4 Genotypic Characterization.................................................................................................................................. 259 24.1.4.1 Molecular Fingerprinting....................................................................................................................... 259 24.1.4.2 PCR and Hybridization Techniques Based on Specific Primers and Probes........................................ 261 24.1.4.3 Sequencing of rRNA and Other Molecular Marker Genes................................................................... 263 24.1.4.4 Temperature and Denaturing Gradient Gel Electrophoresis (TGGE and DGGE)................................ 263 24.2 Methods of the Most Commonly Used Techniques for Lactobacillus Detection............................................................ 264 24.2.1 DNA Sample Preparation..................................................................................................................................... 264 24.2.2 Detection Procedures............................................................................................................................................ 264 24.2.2.1 Identification of Lactobacillus Species by PCR and DNA Sequencing................................................ 264 24.2.2.2 Differentiation of Lactobacillus Species by Multiplex or Simplex PCR............................................... 264 24.2.2.3 Detection and Identification of Lactobacillus Species by DGGE......................................................... 264 24.2.2.4 Quantification of Lactobacillus by Real-Time PCR (SYBR Green)..................................................... 265 24.3 Conclusions....................................................................................................................................................................... 265 Acknowledgments...................................................................................................................................................................... 265 References.................................................................................................................................................................................. 265

24.1  INTRODUCTION The genus Lactobacillus belongs to the lactic acid bacteria (LAB) group, which comprises a broadly defined grampositive bacterial species of nonspore-forming rods or cocobacilli, and which is characterized by the formation of lactic acid as the sole or main end product of carbohydrate metabolism. They are catalase-negative, strictly fermentative, aerotolerant or anaerobic, aciduric or acidophilic, and have complex nutritional requirements and a low GC (guanine + cytosine) content (85%) to lactic acid by the Embden–Meyerhof–Parnas (EMP) pathway, but not pentoses and gluconate because they lack phosphoketolase activity; (ii) facultative heterofermentative lactobacilli that degrade hexoses to lactic acid by the EMP pathway and also ferment pentoses (and often gluconate) because they possess both aldolase and phosphoketolase activities; and (iii) obligately heterofermentative lactobacilli that degrade hexoses by the phosphogluconate pathway producing lactate, ethanol or acetic acid, and carbon dioxide in equimolar amounts; moreover, they ferment pentoses by the same pathway. However, the division of lactobacilli according to the type of fermentation is not in accordance with their phylogenetic clustering as revealed by analysis of their 16S rRNA. The application of polyphasic approaches, which consist of integrating different types of data (phenotypic, genotypic, and phylogenetic) of microorganisms, has been shown to be very useful for taxonomy.7 The use of polyphasic taxonomy has led to taxonomic changes within the Lactobacillus genus. For example, it has led to the reclassification of species, such as L. sobrius strain DSM 16698T as L. amylovorus;8 and genera, such as Pediococcus dextrinicus as L. dextrinicus.9 It has also led to the identification of many new species (e.g., species isolated from sunki, a traditional Japanese pickle: L. kisonensis, L. otakiensis, L. rapi, and L. sunkii).10 At the time of writing (August 2009), in the Approved List of Bacterial Names11 the genus Lactobacillus includes 133 validly described species and 18 subspecies, which reflects its complexity. The description of a large number of new species and the following phylogenetic reexamination of the genus has caused some confusion recently, and earlier lists of identified lactobacilli will continue being modified.12

Molecular Detection of Human Bacterial Pathogens

24.1.2  Ecology Lactobacilli are found in environments where carbohydrates are available, such as food (dairy products, fermented meat, sourdough, vegetables, fruits, and beverages); sewage and plant material; and the respiratory, gastrointestinal, and genital tracts of humans and animals. Lactobacilli are important in the production of foods that involve lactic acid fermentation, notably dairy products (e.g., yogurt and cheese), fermented vegetables (e.g., olives, pickles and sauerkraut), fermented meats (sausages), and sourdough bread. The use of lactobacilli as starter cultures in the food industry has a long history.13 Starter cultures play critical roles in the fermentation processes, development of flavor and texture, and preservation of different fermented foods. For instance, the expression of 15 genes and/or operons was induced in L. sakei during raw-sausage fermentation, reflecting the functional adaptation and roles of this microorganism to the fermentation sausage environment.14 Some Lactobacillus species can also contribute to food spoilage, which are cause of concern in the food industry. The overgrowth of nonstarter species can cause acidification, smear, and out flavors in different products, such as cooked ham and cheeses.15,16 Lactobacillus species are normal residents of the gastrointestinal tract of humans and animals, and have received considerable attention for their possible use as probiotics.17,18 Several recent studies have highlighted the benefits and limitations of the use of specific Lactobacillus strains in different health-related areas, including maintenance of Crohn’s disease remission,19,20 stimulation of the immune system,21,22 prevention of allergic diseases,23 and reduction of the recurrence of Clostridium difficile infections.18,24 For the time being, dairy products are the main vehicle used to supply humans with probiotic strains and, especially, fermented milks, which allow their growth and survival better than other food matrixes. In general, Lactobacillus are avirulent or show low virulence potential; however, members of this genus have been associated with infections in immunosuppressed subjects and patients with health complications, such as bacteremia, endocarditis, and vascular graft infection.25–27 Lactobacilli have also been isolated from carious dentin lesions and are associated with the development of dental caries due to their aciduric characteristics.28

24.1.3  Phenotypic Characterization The phenotypic characterization of lactobacilli usually includes evaluation of morphology, mode of glucose fermentation, growth at different temperatures, configuration of the produced lactic acid, type of fermented carbohydrates, production of methyl esters from fatty acids, and the pattern of whole-cell or cell-wall proteins. However, the use of these traditional methods for the identification of lactobacilli is timeconsuming, requires cultivation, and may lead to erroneous results. In general, phenotypic tests show poor reproducibility

Lactobacillus

and ambiguous results. The expression of phenotypic features depends on the environmental conditions contributing to the variability of the results; for example, growth at different temperatures changes the lipid composition of the membrane of L. acidophilus CRL 640.29 Moreover, lactobacilli share many biochemical and phenotypical characteristics within species of the same genus and other related genera. Altogether, these drawbacks adversely affect the reliability of phenotype-based methods as unique tools for identification of lactobacilli at genus and at species level. Commercial kits based on biochemical and enzymatic analysis has been used for long time as simple and fast methods for the identification of Lactobacillus at species level. Nevertheless, the results of these analyses, for instance using the API 50 CH strips, did not have good reproducibility, probably due to their susceptibility to different environmental and physiological conditions. These data suggest that the use of the current API 50 CH database and other similar methods for identification of Lactobacillus species could lead to misidentification, when applied as the only identification tool.30 Anyway, phenotypic characterization is important for a preliminary classification as well as to get basic knowledge of the properties of Lactobacillus strains. Salt and pH tolerance, growth at certain temperatures, configuration of the produced lactic acid, antibiotic resistance, and bacteriocin production are important characteristics for selecting Lactobacillus strains as possible starter or protective cultures and as probiotics.

24.1.4  Genotypic Characterization Many different genotyping techniques can be applied for species identification, strain differentiation, and quantification. The major advantages of these DNA-based typing methods lie in their discriminatory power, universal applicability, and culture independency.31 Species and closely related strains with similar phenotypic features can be distinguished by DNA-based molecular fingerprinting techniques, including restriction fragment length polymorphism (RFLP), pulse field gel electrophoresis (PFGE), randomly amplified polymorphic DNA (RAPD), and ribotyping: identification and quantification of species and groups can be achieved by molecular techniques based on the use of specific primers and probes, including conventional polymerase chain reaction (PCR), real-time PCR hybridization techniques, and sequencing of rRNA and other gene markers. Moreover, detection and evolution of species in complex ecosystems can be achieved by denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE). These methodologies and examples of their application to the genus Lactobacillus are described below. 24.1.4.1  Molecular Fingerprinting Many common typing methods have been adapted to the genus Lactobacillus. These techniques are applied to differentiate between species and strains, which may belong to the same species, previously isolated on a culture medium. The

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techniques used for the fingerprinting of Lactobacillus species are summarized in Figure 24.1. Restriction Fragment Length Polymorphism. In RFLP analysis, the DNA is digested by restriction enzymes and the resulting restriction fragments are separated according to their lengths by gel electrophoresis. The banding patterns that result are referred as DNA fingerprinting. Because of the high specificity of restriction enzymes and the stability of chromosomal DNA, a reproducible pattern of fragments is obtained after the complete digestion of the chromosomal DNA by a particular enzyme. One general criticism about this method is the complexity of banding patterns. However, this technique can be applied on a selected genetic marker by its amplification with specific primers (PCR–RFLP), reducing its complexity. Then, the generated DNA fingerprinting and the discrimination power between different bacterial species or subspecies, will be dependent on the restriction enzymes used and the length of the amplified fragment. Amplified ribosomal DNA restriction analysis (ARDRA) is essentially PCR–RFLP analysis of the amplified 16S rRNA gene. ARDRA patterns are highly reproducible and comparable between laboratories, but the discriminatory power of this technique is generally low because of the conserved nature of this gene. Other highly conserved genes, such as rpoB, hsp60, or the internal spacer region between the 16S and 23S rRNA genes (ISR), and their restriction by different enzymes have been used for Lactobacillus identification by PCR–RFPL. Chenoll et al. (2006)32 identified L. tucceti as a novel specie based on their singular ISR-DdeI and ISRHaeIII profiles that allowed its differentiation from 68 lactic acid bacteria reference strains and showed that the discriminatory power of the ISR digestion was better than that of the 16S rRNA gene.33 Furthermore, RFLP analysis of the rpoB sequence amplified by PCR has successfully been used to differentiate 12 lactobacilli at special level after two or three digestions (AciI, HinfI, and MseI).32 Pulse Field Gel Electrophoresis. PFGE employs an alternating field of electrophoresis to allow the separation of large DNA fragments obtained from restriction digests of genomic DNA with rare-cutting enzymes (i.e., SfiI, NotI, ApaI, SgrA, and SmaI). PFGE is performed as a standard gel electrophoresis, but the voltage is periodically switched among three directions; one that runs through the central axis of the gel, and two that run at an angle of 120 degrees either side.34 The technique can be more time consuming than other fingerprinting strategies (4–5 days). However, the profile generated by PFGE has a very good discriminatory power. Indeed, PFGE technique has been successfully used for Lactobacillus species and subspecies differentiation.34–36 For example, the diversity and dynamics of Lactobacillus populations were monitored throughout the manufacturing of Camembert cheese. L. paracasei was the species most frequently isolated and these isolates (39) were classified by PFGE into 21 different profiles among the same species.37 The typing efficacy of this technique can be increased by the use of several single restriction enzymes, followed by numerical analysis of the combined patterns. Sánchez et al (2004).35

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Molecular Detection of Human Bacterial Pathogens

(a) RFLP

1

2

3

4

2 3 1

DNA

Restricted DNA with endonucleases

4 Agarose gel A+

(b) PFGE

B–

B+ Cells embedded in agarose

Enzimatic purification of DNA

A– Size separate DNA fragments

(c) RAPD 1

3 4

2

3

1

4 Targeted DNA

2 Agarose gel

(d) Ribotyping

DNA

Restricted DNA with endonucleases

Size separate DNA fragments

Hybridized whith rDNA probes

FIGURE 24.1  Fingerprinting techniques for Lactobacillus species and strains (a) Restriction fragment length polymorphism (RFLP). The DNA fragment is restricted into pieces (for example 1–4) by a restriction enzyme. The restricted DNA is then loaded into a well in an agarose gel and the fragments are separated by electrophoresis, based on fragment size. (b) Pulse field gel electrophoresis (PFGE). Whole cells are embedded in agarose and DNA is enzymatically purified and restricted by rare-cutting enzymes in situ. The agarose plug is then inserted into a well in an agarose gel and the large restricted fragments are separated by an electric current which pulses from different angles. (c) Random amplification of polymorphic DNA (RAPD). PCR is run with arbitrary sequence primers that amplify segments of DNA of unknown identity. The amplified DNA is then loaded into an agarose gel and the fragments are separated by electrophoresis, based on fragment size. (d) Ribotyping. DNA is isolated from a cultured isolate, restricted and size separated in an agarose gel. The gel is then hybridized with a labeled rRNA/rDNA probe, which binds to fragments containing copies of the specific rRNA. Following probe detection, fragments with bound probe are visualized.

used five rare-cutting enzymes (SfiI, NotI, ApaI, SgrA, and SmaI) for PFGE analysis of 149 lactobacilli isolated from fermented vegetables. The use of SfiI and SmaI led to the most informative digestion patterns, and their combined numerical analysis improved the discriminatory power of the technique.35 Some comparative studies of different typing techniques have indicated that isolates that are indistinguishable by PFGE are unlikely to demonstrate substantial differences by other typing techniques.38,39 Nevertheless, PFGE patterns can be used for bacterial identification only when a set of known strains is included in the same analysis and when a suitable and objective method of analysis of the patterns is applied.40

Random Amplification of Polymorphic DNA Finger­ printing. The RAPD technique is a PCR-based discrimination method in which short arbitrary primers (8–12 nucleotides) anneal to multiple random target sequences of the genomic DNA, resulting in patterns of diagnostic value. Low-stringency annealing conditions are used in PCR reactions, which results in the amplification of randomly sized DNA fragments. RAPD–PCR is a molecular-typing method that is very suitable for identification and differentiation of Lactobacillus strains. It is less time consuming than PFGE, but the construction of a database is nearly impossible due to the high variability of the results. If an unidentified strain has

Lactobacillus

to be investigated a set of known strains has to be included in the same RAPD procedure. As the reproducibility of RAPD patterns is occasionally poor, this method needs to be performed under carefully controlled conditions. RAPD technique has been reported as a very useful tool for Lactobacillus differentiation at strain level,15,16,41,42 but other complementary tools are required for species identification, such as sequencing of molecular marker genes, DGGE, or TGGE. In this context, for example, Dimitonova et al. (2008), compared the RAPD patterns of 20 vaginal lactobacilli isolates, generated with the M13V primer. Two main clusters were generated and divided into subgroups. Identical RAPD patterns were shown in different isolates, suggesting that they were multiple isolates from a single species and identical strains. ARDRA-HaeIII analysis and 16S rRNA gene sequencing allowed the identification at species level of seven strains, three as L. fermentum, two as L. gasseri, one as L. brevis, and one as L. salivarius.43 Ribotyping. A ribotype is essentially a RFLP profile consisting of the restriction fragments from a particular genome, which contain rRNA genes. The bacterial chromosomal DNA is totally restricted into multiple fragments, using a specific restriction enzyme. The restricted fragments are then separated by agarose gel electrophoresis and hybridized with 16S and 23S rRNA probes. The hybridization can be carried out directly in the gel using gel hybridization techniques or on a nylon or nitrocellulose membrane following Southern transfer DNA from the gel to the membrane.44 Following probe detection, restriction bands containing copies of the rRNA genes are visualized and the pattern of the band sizes represents a characteristic fingerprint. Ribotyping fingerprint patterns are more stable and more easily interpretable than those obtained by only restriction enzyme analysis; furthermore, they are highly reproducible, and a single rRNA probe can be used for typing all bacteria because of the similarity of ribosomal genes.45 Ribotyping technique also allows the differentiation of Lactobacillus isolates at strain level. In this context, for example, the probiotic strain L. rhamnosus 35 was characterized at molecular level and compared with seven reference strains from the L. casei group by ribotyping.46 This molecular technique is also a useful tool for the identification of new species, for instance, Lactobacillus similis.47 Automated ribotyping has also been used for the characterization of Lactobacillus. For instance, 91 type and reference stains were classified in different ribotypes. Strains of the L. casei group were divided into five ribotyping genotypes, and strains of the L. acidophilus group were divided into two major clusters, one containing L. acidophilus, L. amylovorus, L. crispatus and L. gallinarum, and another containing L. gasseri and L. johnsonii.48 Several studies have compared the discriminatory power of different molecular-typing methods.35,49,50 In general, the most commonly applied techniques can be placed in the following order with respect to their discriminatory power: PFGE > ribotyping > RAPD > ARDRA. The choice of the most appropriate typing method also depends on other

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features such as cost, high throughput capacity, and reproducibility of the fingerprints. 24.1.4.2  P  CR and Hybridization Techniques Based on Specific Primers and Probes Specific oligonucleotide probes and primers targeting rRNA or rDNA have been designed for detection and quantification of different species of the Lactobacillus genus using diverse PCR and hybridization techniques. Lactobacilli are phylogenetically heterogeneous, which causes difficulties in the design of a very specific genus probe, but some group-specific probes and primers have been described.51 Frequently, Lactobacillus group-specific primers cover, in addition to Lactobacillus, Leuconostoc, Pediococcus, and Weissella genera.52,53 Speciesspecific probes and primers have also been designed for the identification of Lactobacillus species, but the probe panel is still incomplete, and the detection of some species by these techniques is not possible yet.51 Conventional and Real-Time PCR. Amplification of DNA by PCR is one of the easiest and most rapid ways of detecting and identifying specific bacterial sequences with the use of adequate primers. A summary of primers and conditions designed for the detection of members of the Lactobacillus genus is shown in Table 24.1. The conventional PCR is sufficiently sensitive to detect Lactobacillus at genus57,58 and at specie level.52–56 More recently, multiple PCR approaches have been designed to detect simultaneously different species in a single PCR reaction. For example, it has been possible to simultaneously distinguish L.  plantarum, L. pentosus and L. paraplantarum species by using a multiplex PCR protocol. The size of the amplicons was 318 bp for L. plantarum, 218 bp for L. pentosus, and 107 bp for L. paraplantarum.59 Conventional PCR can be used for semiquantitative analyses by making dilutions of known amounts of DNA almost identical to the target.60 Real-time PCR is a DNA-based technique that monitors the amplification of the target DNA in real time by fluorescence detection. The amount of DNA corresponding to gene copy numbers or bacterial cells in a sample is determined by measuring the cycle number at which the increase in fluorescence (and therefore cDNA) is exponential. Real-time PCR can be used to quantify bacteria from various samples including milk, feces, food, and water. Contemporary, realtime PCR allows the monitoring of the complete amplification and a more accurate quantification. Even though, the determination of bacterial numbers is difficult due to factors affecting the amplification reaction and variation of the copy number and the ribosomal content of cells in different bacteria.61 In spite of these limitations, sets of species-specific 16S rRNA primers and real-time PCR have been used successfully for monitoring and quantifying, for example, the genus Lactobacillus in feces of patients with celiac disease and healthy controls.62,63 Scheirlinck et al. (2009)64 also studied the predominant sourdough LAB species in the production of sourdough bakeries. Real-time PCR assays were developed using species-specific primers targeting the pheS gene of L. plantarum and L. sanfranciscensis to detect, quantify, and monitor them in sourdough and bakery environments.64

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TABLE 24.1 16S rRNA Targeted PCR Primers Designed for the Detection of Lactobacillus Species Specificity L. acidophilus

L. casei

L. casei/paracasei L. casei/paracasei/ rhamnosus/zeae L. crispatus

L. delbrueckii

L. fermentum L. gallinarum L. gasseri L. johnsonii

L. paracasei

L. plantarum L. reuteri L. rhamnosus

L. salivarius

Sequence (5′–3′) GATCGCATGATCAGCTTATA AGTCTCTCAACTCGGCTATG AGCTGAACCAACAGATTCAC ACTACCAGGGTATCTAATCC GCGGACGGGTGAGTAACACG GCTTACGCCATCTTTCAGCCAA GCGGAAATAGTAGTGTGACGATC GCGGAAATAGGTGCATAGGCG CTCAAAGCCGTGACGGTC ACGTGGTGCTAATAATCCTAGTG GTAATGACGTTAGGAAAGCG ACTACCAGGGTATCTAATCC GTGCTTGCACTGAGATTCGACTTA TGCGGTTCTTGGATCTATGCG TCTTGATTTAATTTTG GCACCGAGATTCAACATGG GCGGACGGGTGAGTAACACG GCTTACGCCATCTTTCAGCCAA GGTTCTTGGATYTATGCGGTATTAG ACATGAATCGCATGATTCAAG AACTCGGCTACGCATCATTG AGCGAGCGGAACTAACAGATTTAC GCGGACGGGTGAGTAACACG GCTTACGCCATCTTTCAGCCAA GGRTGATTTGTTGGACGCTAG GCCGCCTTTCAAACTTGAATC GCACCTGATTGATTTTGGTCG GTTGTTCGCATGAACAACGCTTAA CGGTAATGACGCTGGGGAC CAGTTACTACCTCTATCTTTCTTCACTAC GAGCTTGCCTAGATGATTTTA ACTACCAGGGTATCTAATCC CACTAGACGCATGTCTAGAG AGTCTCTCAACTCGGCTAT GTGCTTGCACCGAGATTCAACATG TGCGGTTCTTGGATCTATGCG CCGAGATTCAACATGG ATCATGATTTACATTTGAGTG TGAATTGACGATGGATCACCAGTG TGCTTGCATCTTGATTTAATTTTG GTGCTTGCATCTTGATTTAATTTT TGCGGTTCTTGGATCTATGCG CGAAACTTTCTTACACCGAATGC ATTCACTCGTAAGAAGT

Hybridization Techniques. Hybridization techniques with specific probes are also useful for the detection and quantification of lactobacilli. Of these techniques, the most commonly used are dot blot hybridization and fluorescence in situ hybridization (FISH). Dot blot hybridization is used for the identification of specific nucleic acid sequences or genes with varying degree of homology and for the estimation of relative amounts of nucleic

Target Site

Reference

205–224 328–309 70–89

52 53 53 54 54 53

61–84 185–165 68 86–104 93–112 213–192 196–169

52

65–89 185–205 305–286 92–102 219–199 86–94 160 78–97 470–442

56 52

170–189 296–277 60–83 185–165 69 96 91 81–104 88–111 213–194 67–91 95

52

55 56 52 56 52

56 56 56 55 56 56 53

52 55 55 55 56 52 56 55

acid with known homology. DNA or RNA is extracted from a sample or cultured bacteria, fixed to a nylon or nitrocellulose membrane and hybridized with a labeled oligonucleotide or DNA fragment complementary to the nucleic acids of interest. For example, dot blot hybridization analysis has been used for screening the presence of Lactobacillus reuteri and the gene encoding for the enzyme responsible for reuterin production (gldC).65

Lactobacillus

In FISH, the detection of rRNA sequences within morphologically intact cells is achieved using fluorescently labeled oligonucleotide probes. Special permeabilization procedures may be required to aid passage of the probe into the cells of some bacteria. The multiple rRNA molecules present in bacterial cells capture the probe, the signal is intensified, and the cells can be detected and numerated by using an epifluorescence microscope or flow cytometry.66,67 For example, using fluorescent L. plantarum-specific targeted oligonucleotide probes, the presence of L. plantarum L2 on the jejunum, ileum and colon of rats was demonstrated, after its oral administration.68 The use of FISH coupled with flow cytometry analysis has also demonstrated that Lactobacillus population is significantly reduced in the intestinal microbiota of patients with celiac disease compared with healthy children.69 24.1.4.3  S equencing of rRNA and Other Molecular Marker Genes Relatedness among organisms is estimated through the comparison of molecular sequences, mainly 16S rRNA encoding genes.70 It is based on several major assumptions: rRNA genes are highly conserved because of the fundamental role of ribosomes in the protein biosynthesis that was developed in the early stages of the evolution of microorganisms. Horizontal gene transfer phenomena among organisms have not involved those genes, and the amount of similarity of these sequences between different individuals is representative of the variation of their genomes. The 16S rRNA gene is a well-conserved universal marker, but there are some shortcomings associated with its use. First, the 16S rRNA genes are so well conserved that it has limited resolving power. Second, although the 16S rRNA gene is a universal marker, different bacterial species have different copy numbers of the gene and this leads to an over- and underrepresentation of some bacterial species. In general, if the 16S rRNA gene sequence identity of two microorganisms is lower than 97%, they are poorly related at the genomic level and, therefore, they are considered to belong to different species. If the 16S rRNA gene sequence identity of two microorganisms is higher than 97%, even if they are identical sequences, they have to be considered closely related, and only total DNA–DNA hybridization data and/or analysis of other more discriminative gene sequences are decisive for the identification at the species level. Phylogenetic and alignment analyses are largely preferred over DNA hybridization tests because the latter are time consuming, complex, and expensive. In contrast, identification through sequence analysis is based on specific DNA amplification, sequencing reaction, and comparison with public databases, which are very fast, more reproducible and much less expensive.12 In general, 16S rRNA gene analysis has been used for the identification of different Lactobacillus species,71,72 but the high similarities of the 16S rRNA gene in this bacterial group usually require the confirmation by the analysis of other markers. Thus, the sequencing of several other genes, such as the recA (encoding recombinase A)42,61,73; tuf gene (encoding

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elongation factor Tu, involved in protein biosynthesis)74,75; hsp60 (groEL, encoding a 60-kDa heat shock protein)76; mal (encoding malolactic enzyme)77; and rpoB (encoding RNA polymerase beta subunit)32 has been used for the identification of lactobacilli. 24.1.4.4  T  emperature and Denaturing Gradient Gel Electrophoresis (TGGE and DGGE) For DGGE and TGGE analysis, RNA or DNA is extracted from an environmental sample and is amplified with specific primers. In DGGE as well as in TGGE, DNA fragments of the same length, but with different sequences, can be separated by electrophoresis through a polyacrylamide gel containing a linear gradient of DNA denaturants formed by urea and formamide (DGGE) or a linear temperature gradient (TGGE). Separation is based on the melting of rRNA genes at specific denaturing points based on their sequence; therefore, each individual sequence will begin to melt at a characteristic denaturing point. The melting changes the conformation of the DNA molecule, slowing its migration though the gel. By using DGGE or TGGE, 50% of the sequence variants can be detected in DNA fragments up to 500 bp. This percentage can be increased to nearly 100% by the attachment of a GC-rich sequence to one side of the DNA fragment. A GC clamp is generally included at the five-end of one of the PCR primers, co-amplified, and thus introduced into the amplified DNA fragments in order to maintain the double-stranded integrity of the amplified products, which improves the detection. The gene sequences from bacterial species in a mixed culture are first amplified using conserved bacterial primers (universal or group-specific primers) that target a hypervariable region, producing amplicons of the same length but with differing sequences that are specific at specie level. DGGE and TGGE allow the separation of these amplicons, producing a molecular fingerprint of the bacterial species present in a sample. The design of primers based on sequence variability in 16S rRNA, 23S rRNA, and other molecular marker genes together with DGGE and TGGE techniques have improved our understanding of the lactobacilli communities present in a variety of complex ecosystems, including the intestinal tract52,78,79 and fermented food.32,42 DGGE and TGGE techniques are useful tools for monitoring both dynamic changes in mixed populations over time and the diversity of Lactobacillus communities because they are rapid and economical. However, they also have several limitations.80 First, only relatively small fragments can be separated by DGGE and TGGE, limiting the amount of sequence information for phylogenetic inferences. Second, the use of different regions of the 16S rRNA and different DGGE and TGGE conditions might result in different resolutions of separation. Finally, DGGE and TGGE only display the DNA fragments obtained from the predominant species present in the community. Several studies revealed that lactobacilli make up less than 1% of the gastrointestinal community, and in this case, the use of a nested approach improves the detection of bands in the denaturing gradient gels. The nested-PCR approach

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consisting in a first amplification with universal bacterial primers and a second amplification with specific primers for Lactobacillus and closely related species.81

24.2  M  ETHODS OF THE MOST COMMONLY USED TECHNIQUES FOR LACTOBACILLUS DETECTION 24.2.1  DNA Sample Preparation Lactobacillus DNA can be extracted with conventional DNA extraction procedures using lysozyme, proteinase K, and sodium dodecyl sulfate, with slight modifications depending on the type of starting material.82 Commercialized kits may also be used for genomic DNA extraction from fresh or frozen human stool or other sample types, using a rapid and simplified procedure. DNA extracts from reference bacterial strains can also be obtained from a high, concentrated cell suspension in 50 µL of sterile water by boiling for 10 min and freezing for 5 min at −20°C. To verify whether a particular DNA extraction method is effective for a specific matrix, it should be tested on samples containing a well-known number of lactobacilli cells.

24.2.2  Detection Procedures 24.2.2.1  I dentification of Lactobacillus Species by PCR and DNA Sequencing Principle.  Lactobacillus spp. can be identified by PCR amplification and sequencing analysis of 16S rRNA using the universal primers 27f and 1492R. The primer sequences are: 27f, 5′-AGAGTTTGATCCTGGCTCAG-3′ 83; and 1495R, 5′-CTACGGCTACCTTGTTACGA-3′.84 Procedure.  The reaction mixture (50-µL) is performed as follows: 5 µL of extracted DNA, 5 µL 10× buffer, 0.2 mM of dNTP, 2.5 mM MgCl2, 10 pmol of each primer, and 1 U of Taq DNA polymerase. Amplification is done using a PCR System programmed as follows: 94°C for 5 min; 30 cycles of 94°C for 1 min, 58°C for 1 min and 72°C for 2 min; and finally, 72°C for 10 min. PCR products (1.5 kb) are electrophoresed on a 1% agarose gel in 1× Tris-Borate-EDTA, and visualize by UV transillumination after ethidium bromide staining. Sequence the PCR products by using the forward primer and the reverse primer. For bacterial identification, partial 16S rRNA gene sequences are compared with sequences deposited in the GenBank database, using the program standard nucleotidenucleotide Basic Local Alignment Search Tool (BLAST). A threshold of 97% similarity in 16S rRNA gene sequence has been proposed for the bacterial-species delineation based on a correlation study between DNA–DNA hybridization results and similarity values for 16S rRNA gene sequences. 24.2.2.2  D  ifferentiation of Lactobacillus Species by Multiplex or Simplex PCR Principle.  The species Lactobacillus plantarum, Lactobacillus pentosus, and Lactobacillus paraplantarum can be

Molecular Detection of Human Bacterial Pathogens

differentiated by doing a unique PCR reaction with recA derived primers. The multiplex PCR assay can be performed with the primers paraF (5′-GTCACAGGCATTACGAAAAC-3′), pentF (5′-CAGTGG CGCGGTTGATATC-3′), planF (5′-CC GTTTATGCGGAACACCTA-3′), and pREV (5′-TCGGG ATTACCAAACATCAC-3′).59 Lactobacillus species can also be detected by simplex PCR using primers specifics for each specie in each reaction, as those summarized in Table 24.1. Procedure.  The PCR mixture (20 µL) is composed of 5 µL of DNA, 2 µL 10× buffer, 1.5 mM MgCl2, 0.12 mM of the primer planF and 0.25 mM of the primers paraF, pentF, and pREV, 0.3 mM of dNTPs, and 1 U of Taq DNA polymerase. PCRs are performed using a PCR System programmed as follows: 94°C for 3 min, 30 cycles of 94°C for 30 s, 56°C for 10 s, and 72°C for 30 s; and final extension at 72°C for 5 min. The PCR products are then separated electrophoretically on an agarose gel and stained with ethidium bromide. L. plantarum, L. pentosus, and L. paraplantarum strains generate a 318-, 218-, and 107-bp fragments, respectively. Similarly, the amplification products of a simple PCR reaction can be detected on an agarose gel and identified according to its expected size by ethidium bromide staining. 24.2.2.3  D  etection and Identification of Lactobacillus Species by DGGE Principle.  Fecal Lactobacillus communities can be monitored by PCR amplification and DGGE analysis of using 16S rRNA group specific primers. Procedure.  Genomic DNA is amplified using the primers Lac1 (5′-AGCAGTAGGGAATCTTCCA-3′); and Lac2 (5′-ATTYCACCGCTACACATG-3′). A GC clamp (5′-CGCCCGGGGCGCGCCCCGGGCGGCCCGGGG GCACCGGGGG-3′) was attached to the reverse primer (Lac2) to obtain PCR fragments suitable for DGGE analysis.52 Amplification can be carried out using a GeneAmp 2400 Thermocycler (Perkin-Elmer). The reaction mixture (50 μL) contained 25-pmol amounts of each primer, 0.2 mM of each dNTP, reaction buffer, 2 mM MgCl2, 25 μg of bovine serum albumin, 2.5 U of Taq polymerase (Amersham Pharmacia Biotech), and 1 μL of DNA solution. The amplification program is 94°C for 2 min; 35 cycles of 94°C for 30 s, 61°C for 1 min, and 68°C for 1 min; and finally, 68°C for 7 min. DGGE analysis of PCR amplicons is carried out on the Dcode Universal Mutation Detection System (Bio-Rad, Richmond, CA). The denaturing gradient of urea and formamide used for PCR products separation is 25%–45%. A 100% denaturant corresponds to 7M urea and 40% (v/v) formamide. DGGE ladders for the identification of Lactobacillus-like populations are prepared by mixing equal amounts of amplicons obtained from selected reference Lactobacillus species using primers Lac1 and Lac2-GC. Selected unknown DGGE bands can be excised from the acrylamide gels and reamplified with primers Lac1 and Lac2. After purification, sequencing is caring by DNA sequencing using an ABI PRISM-3130XL Genetic Analyzer (Applied

265

Lactobacillus

Biosystems, California, USA). Alternatively isolated bands can be cloned into a vector, checked by PCR, and sequenced afterwards. Sequences identities are determined in GenBank database. 24.2.2.4  Q  uantification of Lactobacillus by Real-Time PCR (SYBR Green) Principle.  Genomic DNA has been used to detect and quantify Lactobacillus spp. with high sensibility by using group-specific primers and real-time PCR (SYBR Green). The primer­ ssequences used are: 5′-AGCAGTAGGGAATCTTCCA-3′, and 5′-CACCGCTACACATGGAG-3′.85 Procedure.  Quantitative PCR can be performed with an iCycler iQ apparatus (Bio-Rad, USA) associated with the iCycler optical system interface software (version 2.3; Bio-Rad) or other equivalent equipments. All PCRs are performed in a volume of 25 µL, using 96-well optical grade PCR plates and an optical sealing tape (Bio-Rad). Reaction mixtures for the optimized SYBR Green assays consist of 1:75,000 dilution of SYBR Green (Molecular Probes); 10 mM Tris-HCl (pH 8.8); 150 mM KCl; 0.1% Triton X-100; 2 mM MgCl2; 100 µM each dNTP; 0.5 µM of each primer and 0.02 U Dynazyme II µL−1 (Finnzymes, Finland); and either 5 µL template or water (no-template control). The thermal cycling conditions are an initial DNA denaturation step at 95°C for 5 min, followed by 35 cycles of denaturation at 95°C for 15 s; primer annealing at 58°C for 20 s; extension at 72°C for 30 s, and an additional incubation step at 80°C–85°C for 30 s to measure the SYBR Green fluorescence. Finally, melt curve analysis was performed by slowly cooling the PCRs from 95°C to 60°C (0.3°C per cycle) with simultaneous measurement of the SYBR Green signal intensity. The bacterial concentration from each sample is calculated by comparing the Ct (cycle threshold) values obtained from standard curves. Standard curves can be created by using serial tenfold dilution of pure culture DNA corresponding to 102–109 cells.

24.3  CONCLUSIONS This decade has seen the emergence of numerous molecular approaches for the analysis of diverse aspects of Lactobacillus populations in complex ecosystems, including the gastrointestinal tract and different foods, and for the identification of new species. Quantitative and qualitative detection can be optimally achieved by the combined use of different methods that complement each other. Thus, numbers can be quantified by using labeled group-specific probes and fluorescent in situ hybridization (FISH), coupled with either microscopy or flow cytometry detection, or by real-time PCR. The species diversity can be subsequently determined by PCR in combination with DGGE, TGGE or DNA sequencing. Strain differentiation could be possible by using PFGE, ribotyping, and RAPD. The application of these techniques is contributing to a comprehensive analysis of the ecological distribution, functional roles, and adaptation of Lactobacillus strains to different

environments. In the near future, it can be anticipated that the increasing use of massive sequencing approaches to the analysis of whole bacterial genomes, metagenomics, and transcripomics will lead to the identification of novel species and roles of Lactobacillus strains in human health and food safety and technology.

ACKNOWLEDGMENTS This work was supported by grants AGL2007–66126-C03–01/ ALI and Consolider Fun-C-Food CSD2007–00063 from the Spanish Ministry of Science and Education (MEC). The scholarship to E. Sánchez from Institute Danone is fully acknowledged.

REFERENCES











1. Hammes, W.P., and Vogel, R.F., The genus Lactobacillus. In Wood, B.J.B., and Holzapfel, W.H. (Editors), The Lactic Acid Bacteria, vol. 2, p. 19, Blackie Academic and Professional, London, United Kingdom, 1995. 2. Bernardeau, M., Vernoux, J.P., Henri-Dubernet, S., and Gueguen, M., Safety assessment of dairy microorganisms: The Lactobacillus genus, Int. J. Food Microbiol., 126, 278, 2008. 3. Beijerinck, M.W., Sur les ferments lactiques de l’industrie, Arch. Néerland. Sci. Exact. Nat., 6, 212, 1901. 4. Hammes, W.P., and Hertel, C., The Genera Lactobacillus and Carnobacterium. In Dworkin, M. (Editor), The Prokaryotes: An Evolving Electronic Resource for the Microbiological Community, vol. 3.15, Springer-Verlag, New York, 2003. 5. Orla-Jensen, S., The lactic acid bacteria, Mem. Acad. Roy. Sci. Denmark Sec. Sci., 5, 81, 1919. 6. Collins, M.D. et al., The phylogenetic analysis of the genus Lactobacillus and related acid lactic bacteria as determined by reverse transcriptase sequencing of 16Sr RNA, FEMS Microbiol. Lett., 77, 5, 1991. 7. Vandamme, P. et al., Polyphasic taxonomy, a consensus approach to bacterial systematics, Microbiol. Rev., 60, 407, 1996. 8. Jakava-Viljanen, M. et al., Lactobacillus sobrius Konstantinov et al. 2006 is a later synonym of Lactobacillus amylovorus Nakamura 1981, Int. J. Syst. Evol. Microbiol., 58, 910, 2008. 9. Haakensen, M. et al., Reclassification of Pediococcus dextrinicus (Coster and White 1964) back 1978 (Approved Lists 1980) as Lactobacillus dextrinicus comb. nov., and emended description of the genus Lactobacillus, Int. J. Syst. Evol. Microbiol., 59, 615, 2009. 10. Watanabe, K. et al., Lactobacillus kisonensis sp. nov., Lactobacillus otakiensis sp. nov., Lactobacillus rapi sp. nov. and Lactobacillus sunkii sp. nov., heterofermentative species isolated from sunki, a traditional Japanese pickle, Int. J. Syst. Evol. Microbiol., 59, 754, 2009. 11. List of prokaryotic names with standing in nomenclature, www.bacterio.cict.fr/ 12. Felis, G.E., and Dellaglio, F., Taxonomy of Lactobacilli and Bifidobacteria, Curr. Issues Intest. Microbiol., 8, 44, 2007. 13. Jay, J.M., Modern Food Microbiology, 5th ed. Chapman and Hall, New York, NY, 1996. 14. Hufner, E. et al., Identification of Lactobacillus sakei genes induced during meat fermentation and their role in survival and growth, Appl. Environ. Microbiol., 73, 2522, 2007.

266 15. Sánchez, I. et al., Genetic diversity, dynamics, and activity of Lactobacillus community involved in traditional processing of artisanal Manchego cheese, Int. J. Food. Microbiol., 107, 265, 2006. 16. Antonsson, M., Molin, G., and Ardo, Y., Lactobacillus strains isolated from Danbo cheese as adjunct cultures in a cheese model system, Int. J. Food Microbiol., 85, 159, 2003. 17. Lebeer, S., Vanderleyden, J., and De Keersmaecker, S.C., Genes and molecules of lactobacilli supporting probiotic action, Microbiol. Mol. Biol. Rev., 72, 728, 2008. 18. Guarino, A., Lo, V.A., and Canani, R.B., Probiotics as prevention and treatment for diarrhea, Curr. Opin. Gastroenterol., 25, 18, 2009. 19. Scott, A.C., Young, R.F., and Fedorak, P.M., Comparison of GC-MS and FTIR methods for quantifying naphthenic acids in water samples, Chemosphere, 73, 1258, 2008. 20. Geier, M.S., Butler, R.N., and Howarth, G.S., Inflammatory bowel disease: Current insights into pathogenesis and new therapeutic options; probiotics, prebiotics and synbiotics, Int. J. Food Microbiol., 115, 1, 2007. 21. del Giudice, M.M., and Brunese, F.P., Probiotics, prebiotics, and allergy in children: What’s new in the last year? J. Clin. Gastroenterol., 42, 205, 2008. 22. Reid, G., Devillard, E., Probiotics for mother and child, J. Clin. Gastroenterol., 38, 94, 2004. 23. Isolauri, E., and Salminen, S., Probiotics: Use in allergic disorders: A Nutrition, Allergy, Mucosal Immunology, and Intestinal Microbiota (NAMI) Research Group Report, J. Clin. Gastroenterol., 42, 91, 2008. 24. Sazawal, S. et al., Efficacy of probiotics in prevention of acute diarrhoea: A meta-analysis of masked, randomised, placebo-controlled trials, Lancet Infect. Dis., 6, 374, 2006. 25. Svec, P. et al., Identification of lactic acid bacteria isolated from human blood cultures, FEMS Immunol. Med. Microbiol., 49, 192, 2007. 26. Chazan, B. et al., Bacteremia and pyelonephritis caused by Lactobacillus jensenii in a patient with urolithiasis, Isr. Med. Assoc. J., 10, 164, 2008. 27. Tommasi, C. et al., Diagnostic difficulties of Lactobacillus casei bacteraemia in immunocompetent patients: A case report, J. Med. Case Rep., 2, 315, 2008. 28. Teanpaisan, R., and Dahlen, G., Use of polymerase chain reaction techniques and sodium dodecyl sulfate-polyacrylamide gel electrophoresis for differentiation of oral Lactobacillus species, Oral Microbiol. Immunol., 21, 79, 2006. 29. Murga, M.L. et al., Influence of growth temperature on cryotolerance and lipid composition of Lactobacillus acidophilus, J. Appl. Microbiol., 88, 342, 2000. 30. Boyd, M.A., Antonio, M.A., and Hillier S.L., Comparison of API 50 CH strips to whole-chromosomal DNA probes for identification of Lactobacillus species, J. Clin. Microbiol., 43, 5309, 2005. 31. Farber, J.M., An introduction to the hows and whys of molecular typing, J. Food Prot., 59, 1091, 1996. 32. Claisse, O., Renouf, V., and Lonvaud-Funel, A., Differen­ tiation of wine lactic acid bacteria species based on RFLP analysis of a partial sequence of rpoB gene, J.  Microbiol. Methods, 69, 387, 2007. 33. Chenoll, E., Macián, M.C., and Aznar R., Lactobacillus rennini sp. nov., isolated from rennin and associated with cheese spoilage, Int. J. Syst. Evol. Microbiol., 56, 449, 2006. 34. Klein, G. et al., Taxonomy and physiology of probiotic lactic acid bacteria, Int. J. Food Microbiol., 41, 103, 1998.

Molecular Detection of Human Bacterial Pathogens 35. Sánchez, I., Sesena, S., and Palop, LL., Polyphasic study of the genetic diversity of lactobacilli associated with “Almagro” eggplants spontaneous fermentation, based on combined numerical analysis of randomly amplified polymorphic DNA and pulsed-field gel electrophoresis patterns, J. Appl. Microbiol., 97, 446, 2004. 36. Roy, D. et al., Molecular identification of potentially probiotic lactobacilli, Curr. Microbiol., 40, 40, 2000. 37. McLeod, A. et al., Diversity of Lactobacillus sakei strains investigated by phenotypic and genotypic methods, Syst. Appl. Microbiol., 31, 393, 2008. 38. Gordillo, M.E., Singh, K.V., and Murray, B.E., Comparison of ribotyping and pulsed-field gel electrophoresis for subspecies differentiation of strains of Enterococcus faecalis, J. Clin. Microbiol., 31, 1570, 1993. 39. Olsen, J.E. et al., Clonal lines of Salmonella enterica serotype Enteritidis documented by IS200-, ribo-, pulsed-field gel electrophoresis and RFLP typing, J. Med. Microbiol., 40, 15, 1994. 40. Nigatu, A., Ahrne, S., and Molin, G., Randomly amplified polymorphic DNA (RAPD) profiles for the distinction of Lactobacillus species, Antonie Van Leeuwenhoek, 79, 1, 2001. 41. Catzeddu, P. et al., Molecular characterization of lactic acid bacteria from sourdough breads produced in Sardinia (Italy) and multivariate statistical analyses of results, Syst. Appl. Microbiol., 29, 138, 2006. 42. Siragusa, S. et al., Taxonomic structure and monitoring of the dominant population of lactic acid bacteria during wheat flour sourdough type I propagation using Lactobacillus sanfranciscensis starters, Appl. Environ. Microbiol., 75, 1099, 2009. 43. Dimitonova, S.P. et al., Phenotypic and molecular identification of lactobacilli isolated from vaginal secretions, J. Microbiol. Immunol. Infect., 41, 469, 2008. 44. Wiedmann, M., Molecular subtyping methods for Listeria monocytogenes, J. AOAC Int., 85, 524, 2002. 45. Mohania, D. et al., Molecular approaches for identification and characterization of lactic acid bacteria, J. Dig. Dis., 9, 190, 2008. 46. Coudeyras, S. et al., Taxonomic and strain-specific identification of the probiotic strain Lactobacillus rhamnosus 35 within the Lactobacillus casei group, Appl. Environ. Microbiol., 74, 2679, 2008. 47. Kitahara, M., Sakamoto, M., and Benno, Y., Lactobacillus similis sp. nov., isolated from fermented cane molasses, Int. J. Syst. Evol. Microbiol., 60, 187, 2009. 48. Ryu, C.S. et al., Characterization of the Lactobacillus casei group and the Lactobacillus acidophilus group by automated ribotyping, Microbiol. Immunol., 45, 271, 2001. 49. Ruiz, P. et al., Intraspecific genetic diversity of lactic acid bacteria from malolactic fermentation of Cencibel wines as derived from combined analysis of RAPD-PCR and PFGE patterns, Food Microbiol., 25, 942, 2008. 50. Tynkkynen, S. et al., Comparison of ribotyping, randomly amplified polymorphic DNA analysis, and pulsed-field gel electrophoresis in typing of Lactobacillus rhamnosus and L. casei strains, Appl. Environ. Microbiol., 65, 3908, 1999. 51. Satokari, R.M. et al., Molecular approaches for the detection and identification of bifidobacteria and lactobacilli in the human gastrointestinal tract, Syst. Appl. Microbiol., 26, 572, 2003. 52. Furet, J.P., Quénée, P., and Tailliez, P., Molecular quantification of lactic acid bacteria in fermented milk products using realtime quantitative PCR, Int. J. Food Microbiol., 97, 197, 2004.

Lactobacillus 53. Walter, J. et al., Detection and identification of gastrointestinal Lactobacillus species by using denaturing gradient gel electrophoresis and species-specific PCR primers, Appl. Environ. Microbiol., 66, 297, 2000. 54. Fujimoto, J. et al., Identification and quantification of Lactobacillus casei strain Shirota in human feces with strainspecific primers derived from randomly amplified polymorphic DNA, Int. J. Food Microbiol., 126, 210, 2008. 55. Chagnaud, P. et al., A. Rapid PCR-based procedure to identify lactic acid bacteria: Application to six common Lactobacillus species, J. Microbiol. Methods, 44, 139, 2001. 56. Byun, R. et al., Quantitative analysis of diverse Lacto­ bacillus species present in advanced dental caries, J. Clin. Microbiol., 42, 3128, 2004. 57. Walter, J. et al., Detection of Lactobacillus, Pediococcus, Leuconostoc, and Weissella species in human feces by using group-specific PCR primers and denaturing gradient gel electrophoresis, Appl. Environ. Microbiol., 67, 2578, 2001. 58. Heilig, H.G. et al., Molecular diversity of Lactobacillus spp. and other lactic acid bacteria in the human intestine as determined by specific amplification of 16S ribosomal DNA, Appl. Environ. Microbiol., 68, 114, 2002. 59. Torriani, S., Felis, G.E., and Dellaglio, F., Differentiation of Lactobacillus plantarum, L. pentosus, and L. paraplantarum by recA gene sequence analysis and multiplex PCR assay with recA gene-derived primers, Appl. Environ. Microbiol., 67, 3450, 2001. 60. Morrison, C., and Gannon, F., The impact of the PCR plateau phase on quantitative PCR, Biochim. Biophys. Acta, 1219, 493, 1994. 61. von, W.F., Gobel, U.B., and Stackebrandt, E., Deter­ mination of microbial diversity in environmental samples: pitfalls of PCR-based rRNA analysis, FEMS Microbiol. Rev., 21, 213, 1997. 62. Collado, M.C. et al., Specific duodenal and faecal bacterial groups associated with paediatric celiac disease, J. Clin. Pathol., 62, 264, 2009. 63. De Palma, G. et al., Effects of a gluten-free diet on gut microbiota and immune function in healthy adult human subjects, Br. J. Nutr., 102, 1154, 2009. 64. Scheirlinck, I. et al., Molecular source tracking of predominant lactic acid bacteria in traditional Belgian sourdoughs and their production environments, J. Appl. Microbiol., 106, 1081, 2009. 65. Cadieux, P. et al., Evaluation of reuterin production in urogenital probiotic Lactobacillus reuteri RC-14, Appl. Environ. Microbiol., 74, 4645, 2008. 66. Amann, R.I. et al., Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations, Appl. Environ. Microbiol., 56, 1919, 1990. 67. Zoetendal, E.G., Akkermans, A.D., and de Vos, W.M., Temperature gradient gel electrophoresis analysis of 16S rRNA from human fecal samples reveals stable and hostspecific communities of active bacteria, Appl. Environ. Microbiol., 64, 3854, 1998. 68. Wang, B. et al., Isolation of adhesive strains and evaluation of the colonization and immune response by Lactobacillus plantarum L2 in the rat gastrointestinal tract, Int. J. Food Microbiol., 132, 59, 2009.

267 69. Nadal, I. et al., Imbalance in the composition of the duodenal microbiota of children with celiac disease, J. Med. Microbiol., 56, 1669, 2007. 70. Woese, C.R., Bacterial evolution, Microbiol. Rev., 51, 221, 1987. 71. Duan, Y. et al., Identification and characterization of Lactic Acid Bacteria isolated from Tibetan Qula cheese, J. Gen. Appl. Microbiol., 54, 51, 2008. 72. Ennahar, S., Cai, Y., and Fujita, Y., Phylogenetic diversity of lactic acid bacteria associated with paddy rice silage as determined by 16S ribosomal DNA analysis, Appl. Environ. Microbiol., 69, 444, 2003. 73. Felis, G.E. et al., Comparative sequence analysis of a recA gene fragment brings new evidence for a change in the taxonomy of the Lactobacillus casei group, Int. J. Syst. Evol. Microbiol., 51, 2113, 2001. 74. Ventura, M. et al., Analysis, characterization, and loci of the tuf genes in lactobacillus and bifidobacterium species and their direct application for species identification, Appl. Environ. Microbiol., 69, 6908, 2003. 75. Chavagnat, F. et al., Comparison of partial tuf gene sequences for the identification of lactobacilli, FEMS Microbiol. Lett., 217, 177, 2002. 76. Blaiotta, G. et al., Lactobacillus strain diversity based on partial hsp60 gene sequences and design of PCR-restriction fragment length polymorphism assays for species identification and differentiation, Appl. Environ. Microbiol.,74, 208, 2008. 77. Groisillier, A., and Lonvaud-Funel, A., Comparison of partial malolactic enzyme gene sequences for phylogenetic analysis of some lactic acid bacteria species and relationships with the malic enzyme, Int. J. Syst. Bacteriol., 49, 1417, 1999. 78. Scanlan, P.D. et al., Culture-independent analyses of temporal variation of the dominant fecal microbiota and targeted bacterial subgroups in Crohn’s disease, J. Clin. Microbiol., 44, 3980, 2006. 79. Sanz, Y. et al., Differences in faecal bacterial communities in celiac and healthy children as detected by PCR and denaturing gradient gel electrophoresis, FEMS Immunol. Med. Microbiol., 51, 562, 2007. 80. Muyzer, G., and Smalla, K., Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology, Antonie Van Leeuwenhoek, 73, 127, 1998. 81. Nielsen, D.S. et al., Case study of the distribution of mucosa-associated Bifidobacterium species, Lactobacillus species, and other lactic acid bacteria in the human colon, Appl. Environ. Microbiol., 69, 7545, 2003. 82. Yu, Z., and Morrison, M., Improved extraction of PCRquality community DNA from digesta and fecal samples, Biotechniques, 36, 808, 2004. 83. Mora, D. et al., Genotypic characterization of thermophilic bacilli: A study on new soil isolates and several reference strains, Res. Microbiol., 149, 711, 1998. 84. 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. 85. Malinen, E. et al., Analysis of the fecal microbiota of irritable bowel syndrome patients and healthy controls with real-time PCR, Am. J. Gastroenterol., 100, 373, 2005.

25 Leuconostoc María Mar Tomás and Germán Bou CONTENTS 25.1 Introduction...................................................................................................................................................................... 269 25.1.1 Taxonomy of Genus Leuconostoc........................................................................................................................ 269 25.1.2 Isolation and Identification of Leuconostoc spp................................................................................................... 269 25.1.3 Virulence and Clinical Significance..................................................................................................................... 272 25.1.4 Antibiotic Susceptibilities..................................................................................................................................... 272 25.1.5 Molecular Identification of Leuconostoc spp....................................................................................................... 272 25.2 Methods............................................................................................................................................................................ 274 25.2.1 Sample Preparation............................................................................................................................................... 274 25.2.2 Detection Procedures............................................................................................................................................ 275 25.3 Conclusion........................................................................................................................................................................ 275 References.................................................................................................................................................................................. 275

25.1  INTRODUCTION Members of the genus Leuconostoc (leucus, clear, light; nostoc, algal generic name; leuconostoc, colorless nostoc) are facultatively anaerobic, catalase-negative, gram-positive cocci organized in pairs and chains.1 They are heterofermentative lactic acid bacteria found naturally in milk, grass, herbage, grapes, many vegetables, meat, and sugar.2,3 Members of this group are used in dairy fermentations to produce aromatic compounds.4

25.1.1  Taxonomy of Genus Leuconostoc Species of Leuconostoc are the only catalase-negative cocci that are vancomycin resistant, PYR and LAP negative, and that produce CO2 from glucose. They could appear morphologically similar to:

1. Genus Weissella, which includes organisms previously classified as Leuconostocs and the species formerly named Lactobacillus confusus. 2. Alpha-hemolytic or nonhemolytic Enterococcus or Lactococcus spp. It happens when these bacteria grown on the surface of 5% sheep blood agar plates. In the past it was known that Leuconostoc spp. was associated with human diseases5 because these bacteria were generally identified as variants of established human pathogens, such as viridans streptococci or enterococci, or more frequently, were reported as unidentified gram-positive cocci. As many as 31% of Leuconostoc spp. react with the group D antiserum and may be reported as



unidentified Enterococcus spp. The nonserogroup D bacteria have characteristics that sometimes lead to their identification as Streptococcus uberis, S. bovis, S. sanguis, S. equinus, S. anginosus group, or Aerococcus-like or Lactobacillus-like bacteria. 3. The heterofermentative vancomycin-­resistant Lactobacillus spp. However, species of Leuconostoc spp. tend to be more coccoid-like than the Lactobacillus spp. Cautious interpretation of the Gram stain coloring from growth of these bacteria in thioglycolate broth is important for differentiation of leuconostocs and gas-forming lactobacilli.

With the recognition of Leuconostoc spp. as human pathogens, such mistakes in identification are now odd because of the unique results produced by leuconostocs in the tests listed in Table 25.1.6

25.1.2  Isolation and Identification of Leuconostoc spp. Isolation Procedure. Generally, there are no special requirements for isolation of the group of bacteria discussed here; general recommendations for the culture of blood, body fluids, and other specimens should be followed. These organisms are likely to be isolated on rich, nonselective media (e.g., blood and chocolate agars and thioglycolate broth) since they are nutritionally fastidious. If selective isolation of members of the vancomycin-resistant genera Leuconostoc and Pediococcus is desired, Thayer–Martin medium may be used to inhibit normal flora or other contaminating microorganisms.34 269

270

Molecular Detection of Human Bacterial Pathogens

TABLE 25.1 Basic Phenotypic Characteristics of Catalase-Negative, Gram-Positive Cocci Organisms (reference[s])

PYR

LAP

NaCl

BE

Abiotrophia defectiva7–9 Aerococcus christensenii10,11 Aerococcus sanguinola11,12 Aerococcus urinae11,13,14 Aerococccus urinaehominis11,15 Aerococcus viridansa,5,11 Dolosicoccus16 Dolosigranulum11,17,18 Enterococcusc,d,5 Facklamia languidae,18,19 Facklamia sppe,18,20–22 Gemella haemolysansf,5 Gemella sppf,5,23,24 Globicatella5,25–27 Granulicatella adiacensg,7–9 Granulicatella elegansg,7–9 Helcococcus kunziia,5,28 Ignavigranum18,26,29 Lactococcusd,h,5 Leuconostoc5 Pediococcus5 Streptococcush,i,5 Tetragenococcus5 Vagococcusc,5 Weissella30,31

+ − + − −

+ + + + −

− + + + +

− − + − −

− + V

+ + + + + + + + V

− − + + + + + + − + + − + + − + + + + ND

+ − + + + + − − + − − + + V V V V

V ND

+b ND

− + ND ND

+ + − − − ND

+ V ND

+ + +

+ + + + V − − V − + V

− − + − − − − + V + V

ESC

+

+ ND ND + − + ND ND V ND + +

MOT

HIP

SAT

ARG

BGUR

− − ND

− + + + +

+ − − − −

− − − − −

− − + + +

− − − V

V ND

− ND

ND ND

− ND

− ND

− − − − − − − − − − − − − − + −

− + ND ND

− − − − − − − − − + + − V

− V ND

− ND ND ND ND

− ND

− ND

+ − + − − ND ND ND ND ND ND ND

− − − − ND − ND

− − − + − − ND ND ND ND ND ND +

+ − − ND ND ND ND ND ND ND ND

MORPH

VAN

Chains Clusters Clusters Clusters Clusters

S S ND S ND

Clusters Chains Clusters Chains Clusters Chains Clusters Chains Chains Chains Chains Clusters Chains Chains Chains Clusters Chains Clusters Chains Short rods coccobacilli in pairs and chains

S S S S/R S S S S S S S S S S R R S S S R

Source: From Murray, P.R., Baron, E.J., Jorgensen, J.H., Landry, M.L., and Pfaller, M.A. (Editors), Manual of Clinical Microbiology, Vol. 1, 31: 443–454, ASM PRESS, Washington, DC, 2007. With permission from American Society of Microbiology. Abbreviations and symbols: PYR, production of pyrrolidonyl arylamidase; LAP, production of leucine aminopeptidase; NaCl, growth in 6.5%NaCl; BE, hydrolysis of esculin in the presence of 40% bile; ESC, hydrolysis of esculin; MOT, motility; HIP, hydrolysis of hippurate; SAT, satelliting behavior; ARG, hydrolysis of arginine; BGUR, production of β-glucuronidase; MORPH, cellular arrangement; VAN, susceptibility to vancomycin; +, ≥90% of strains positive; -, ≤10% of strains positive; V, variable (60%–90% of strains positive); ND, no data; chains, cells arranged primarily in pairs and chains; clusters, cells arranged primarily in clusters, tetrads, or irregular groups; S, susceptible; R, resistant. a Although H. hunzii shares some phenotypic traits with A. viridans, H. kunzii is facultative and usually nonhemolytic, in contrast to A. viridans, which favors an anaerobic growth atmosphere and is alpha-hemolytic. Two additional species of Helcococcus (H. sueciensis and “H. pyogenes”) have been proposed32,33 both based on the isolation of single strains. These new species display negative reactions in the PYR test, in contrast to H. kunzii. b LaClaire and Facklam18 note that strains of Dolosigranulum are esculin hydrolysis positive when tested on conventional media, but the original description of this organism notes a lack of esculin hydrolysis activity. c Most enterococcal strains are capable of growth at 45°C, which differentiates them from vagococci have been reported as testing positive with a commer  cially available nucleic acid probe for members of the genus Enterococcus. d Phenotypically similar strains of enterococci and lactococci can be differentiated with a commercially available nucleic acid probe for members of the genus Enterococcus. e F. languida cells are characteristically arranged in clusters, and cells of other Facklamia species are arranged in pairs and chains. f G. haemolysans cells are arranged in pairs and clusters, and the cells of other Gemella species are arranged in pairs and chains.   g Formerly classified as a member of the genus Abiotrophia. h Most lactococcal strains are capable of growth at 10°C, which differentiates them from streptococci, which may be phenotypically similar. i Streptococcus pyogenes and some strains of S. pneumoniae are PYR positive.

271

Leuconostoc

Attempts have been made to develop media for the isolation and enumeration of these organisms, and both selective and differential media have been described; however, no medium has proven satisfactory. Comprehensive reviews of differential and selective media for Leuconostoc have been published by Garvie,35 Teuber and Geis,36 and Cogan.37 Most media are based on the ability of leuconostocs to utilize citrate, which is recognized by the presence of halos around colonies growing on media containing insoluble calcium citrate.38 Such differentiation is not accurate since not all leuconostocs utilize citrate; furthermore, other bacteria associated with green plants, such as Lactobacillus species39 and Lactococcus lactis subsp. Lactis biovar. diacetylactis, utilize citrate.40 As Leuconostoc species are resistant to vancomycin and Lactococcus species and some lactobacilli are sensitive to vancomycin, we considered inclusion of this antibiotic in a Leuconostoc selective medium (LUSM medium) for the isolation of Leuconostoc spp. The proposed medium contains 1.0% glucose, 1.0% Bacto Peptone (Difco), 0.5% yeast extract (BBL), 0.5% meat extract (Difco), 0.25% gelatin (Difco), 0.5% calcium lactate, 0.05% sorbic acid, 75 ppm of sodium azide (Sigma), 0.25% sodium acetate, 0.1% (vol/vol) Tween 80, 15% tomato juice, 30 μg of vancomycin (Sigma) per mL, 0.20 μg of tetracycline (Serva) per mL, 0.5 mg of cysteine hydrochloride per mL, and 1.5% agar (Difco). The LUSM medium has been successfully used for the isolation and enumeration of Leuconostoc spp. in dairy products and vegetables.2 Identification of Species. The physiological reactions that are useful for identifying the five Leuconostoc spp. that have been isolated from humans infections are listed in Table 25.1.5 All of the isolates were PYR, Voges-Proskauer, and LAP negative, produced gas from glucose in MRS broth, and were resistant to vancomycin.5 Identification of Leuconostoc species that fails to grow in the laboratory at 35°C to 37°C is not included in Table 25.2 (infections in humans by these bacteria are unlikely). Leuconostoc mesenteroides and L. pseudomesenteroides species are sometimes confused with each other. Prolonged incubation of NaCl broth may be necessary for correct

differentiation between the 6.5% NaCl-tolerant L. mesenteroides and the 6.5% NaCl-intolerant L. pseudomesenteroides. Any increase in the number of bacteria in the NaCl broth signifies that the bacterium is L. mesenteroides. Moreover, L. paramesenteroides is considered a species of Leuconostoc with characteristics similar to those of L. mesenteroides. Dextran production by L. mesenteroides is used to differentiate these species. However, current phylogenetic analysis has indicated that L. paramesenteroides is more closely related to the lactobacilli than to members of the genus Leuconostoc,41,42 and there has been an informal proposal to reclassify this and related species into a new genus, Weisella.42 Morphologically, there are some characteristic differentiates between Lactobacillus spp. and Leuconostoc spp. One of them is that Leuconostoc spp. appear as chains of cocci. Leuconostoc citreum is also clearly distinguishable from the other clinically significant species of Leuconostoc because it is the only one that does not ferment either raffinose or melibiose. As its name implies, this bacterium also produces a characteristic yellow pigment that can be observed in bacteria grown in MRS broth.5 The only clinical isolate of Leuconostoc spp. that does not hydrolyze esculin is Leuconostoc lactis (Table 25.2). As with L. paramesenteroides, this species does not produce dextran from glucose. These characteristics clearly differentiate this species from other Leuconostoc species. Finally, the other features to distinguish among these five species of Leuconostoc are (Table 25.2):

1. Alpha hemolysis on Trypticase soy agar with 5% sheep blood (BBL, Cockeysville, Md.). 2. Susceptibility to bacitracin. 3. Serogroup D antigen.

Methods of identifying Leuconostoc cremoris, L. carnosum, L. oenos, L. fallax, L. argentinum, and L. gelidum have been studied in different works.43,44 A new species, Leuconostoc fallax was investigated by reverse transcriptase sequencing of 16S rRNA in 1991.43

TABLE 25.2 Positive Reactions of Centers for Disease Control and Prevention Clinical Isolates of Leuconostoc spp. % Positive in Given Test Species

ESC

LM

RAF

MEL

NaCl

ARA

TRE

SUCA

L. mesenteroides L. pseudomesenteroides L. paramesenteroides L. citreum L. lactis

100 100 100 100 4

11 54 50 4 37

94 77 100 8 96

100 100 10 0 100

89 0 100 63 48

97 62 100 92 52

97 92 100 100 48

74 67 0 83 15

Source:  From Facklam, R., and Elliott, J.A., Clin. Microbiol. Rev., 8, 479, 1995. With permission from American Society of Microbiology. Abbreviations: ESC, esculin hydrolysis; LM, acid and clot production in litmus milk; NaCl, growth in broth containing 6.5%NaCl; SUCA, dextran production on sucrose agar; Acid formation from; RAF, raffinose; MEL, melibiose; ARA, arabinose; TRE, trehalose.

272

25.1.3  Virulence and Clinical Significance The Center for Disease Control and Prevention (CDC)5 carried out a study of the virulence of Leuconostoc species. For the majority of the isolates studied, there was a lack of information about underlying disease in patients, and the possibility that Leuconostoc spp. are primarily opportunistic pathogens of debilitated individuals could not be confirmed. However, considering the wide distribution of these species in the environment and the relatively few infections that they cause, it is assumed that they are poorly virulent for healthy humans. It has been demonstrated that dextran-producing Leucono­ stoc strains are able to inhibit the formation of Streptococcus mutans biofilm in vitro.45 Moreover, Leuconostoc mesente­ roides and Lactococcus lactis have been used as probiotic strains for Aeromonas salmonicida infection in brown trout.46 Handweger and colleagues47 noted host defense impairment, invasive procedures breaching the integument, gastrointestinal symptoms, and prior antibiotic treatment as common features among adult patients infected with Leuconostoc spp. The clinical significance of infection with these bacteria in immunocompetent patients is not clear. For example, in one study the bacteria were eliminated by several patients without the use of antimicrobial agents.48 However, a clinic case involving pulmonary abscess caused by Leuconostoc species in an immunocompetent patient has been reported recently.49 Infections in immunocompromised patients can be severe, and identification of the bacterium as a Leuconostoc spp. is essential to ensure that an appropriate antimicrobial agent is used to treat the patient.50 Invasive procedures recognized as risk factors associated with this infection include total parenteral nutrition through a central venous catheter, and gastrostomy for enteral feeding.51–53 Gastrointestinal symptoms identified as risk factors include short gut syndrome. Several studies have described a link between short bowel syndrome and Leuconostoc bacteremia in pediatric patients.51–53 Finally, prior administration of antibiotics, particularly vancomycin, has been implicated in infections by Leuconostoc. The bacteria are often associated with a polymicrobial infection and are isolated after the patient has been treated with vancomycin to eliminate the suspected pathogen.47,54–56 However, a review of Leuconostoc bacteremia in pediatric patients52 concluded that in the cases of pediatric patients with short bowel syndrome, prior administration of vancomycin was not a required risk factor for development of this infection. Moreover, predisposition to Leuconostoc bacteremia has been observed among neonates, suggesting that infants may become colonized during delivery, by Leuconostoc spp. inhabiting the maternal genital tract. Leuconostoc spp. have occasionally been isolated from humans in cases of bacteremia or septicemia,5,48,50,57 meningitis,58,59 and peritonitis. Some reports have implicated leuconostocs as agents of infection in abdominal abscess,60 osteomyelitis,61 ventriculitis,62 postsurgical endophthalmitis,63 hepatic dysfunction,53 and pneumonia.49 An outbreak of

Molecular Detection of Human Bacterial Pathogens

nosocomial urinary tract infection due to Leuconostoc mesenteroides was reported at a tertiary care centre in northern India.64 These species have not previously been considered as agents that cause severe hospital outbreaks threatening the lives of large numbers of persons. However, three previous reports have described hospital transmission of Leuconostoc spp. Two of these affected a small number of patients, and no epidemiological studies were conducted to clarify the genetic relationship among the bacterial strains involved or the source of the nosocomial infection. The third outbreak occurred in Spain, in 2008,50 and was the largest nosocomial outbreak caused by Leuconostoc spp. worldwide. In this latest example, parenteral nutrition was suggested as the most likely source of infection. Treatment with high-dose penicillin or clindamycin47,54,55,58 or with tobramycin55 has been reported to be successful. Swenson et al.65 found that none of the isolates were resistant to imipenem, minocycline, chloramphenicol, gentamicin, or daptomycin. Daptomycin has also been used to treat infections caused by Leuconostoc spp.66

25.1.4  Antibiotic Susceptibilities The lack of standardized methods and interpretation criteria for antibiotic susceptibility testing led Swenson et al.65 to carry out a study of susceptibility in Leuconostoc mesenteroides, L. paramesenteroides, L. citreum, L. pseudomesenteroides, L. lactis, and L. dextranicum with 79 samples (Table 25.3). However, information on the in vitro antimicrobial susceptibility of L. garvieae is scarce. Glucopeptides. Members of the vancomycin-resistant genera Leuconostoc and Pediococcus are also resistant to teicoplanin. Moreover, most strains are susceptible to daptomycin.65,67 𝛃-lactam Agents. MICs of ampicillin were high, but MICs of penicillin were in the range of 0.25–2.0 µg/mL in 96% of the isolates tested.65 Among cephalosporins, cephalothin was overall the most active cephalosporin tested, but cefuroxime, ceftizoxime, and cefotaxime were more active against strains of L. citreum. Except for strains of L. mesenteroides and L. pseudomesenteroides, the other Leuconostoc spp. are usually susceptible to impenem. To date, no β-lactamase has been detected in any of the strains. Macrolides. All strains of Leuconostoc were susceptible to erythromycin, roxithromycin, and josamycin. Clindamycin MICs were higher (0.12–2 µg/mL) in some strains but these samples did not display greater resistance to erythromycin. Aminoglycosides. Gentamicin was the most active of the four aminoglycosides tested and displayed the greatest activity against leuconostocs.

25.1.5  Molecular Identification of Leuconostoc spp. PCR-Based Methods. In 2000, Lee et al.68 developed a multiplex PCR-based method for rapid and reliable identification of Leuconostoc species, by using species-specific primers

0.5–1 0.12–0.5 0.12–0.5 0.5 0.03 0.5 0.25–2

Leuconostoc mesenteroides (19) L. citreum (12) L. pseudomesenteroides (4) L. lactis (3)

L. dextranicum (1) L. paramesenteroides (1) Leuconostoc species (7)

0.12 32 1–32

2–16 0.5–2 0.5–4 2–4

Cephalothin

1 4 2–128

≤0.25–4 1–16 4–16

≤0.25–1 0.5–16 2–8 0.5 8 0.5–16

8–64

Ceftriaxone

8–32

Cefuroxime

≤0.06 0.12–4

0.12

2–8 1–2 1–8 2–4

Imipenem

Range of MICs (µg/mL) Rifampin 0.5–8 1–2 1–2 1–8 0.06 64 1–64

Erythromycin ≤0.015–0.06 0.03–0.06 0.06 0.06 ≤0.015 0.06 0.06

Source:  Swenson, J.M., et al., Antimicrob. Agents Chemother., 34, 543, 1990. With permission from American Society of Microbiology.

Penicillin

Organism (n)

TABLE 25.3 Range of MICs by Species for Selected Antimicrobial Agents

≤0.25 ≤0.25 ≤0.25–0.5

≤0.25–0.5 ≤0.25 ≤0.25–0.5 ≤0.25–0.5

Gentamicin

1 2 1–4

1–4 0.5–2 2–4 2

Ciprofloxacin

Leuconostoc 273

274

targeted to the genes encoding 16S rRNA. This assay allowed the detection and differentiation of Leuconostoc species in mixed populations from natural sources, as well as from pure cultures, within 3 hours. Arbitrarily primed (AP)-PCR can be used to generate typical DNA band profiles. A single triplicate AP-PCR (TAP-PCR) procedure with an 18-mer oligonucleotide, which enabled fingerprinting of major genera of lactic acid bacteria at the strain level, was developed in 2000.69 The authors used this assay to discriminate Leuconostoc cremoris from other lactic acid bacteria, such as Lactobacillus caseii, Bifidobacterium breve, Lactobacillus plantarum, Pediococcus pentosaceus, and P. cerevisiae. Moschetti et al.70 also used randomly amplified polymorphic DNA analysis with a 10-mer oligonucleotide to characterize Leuconostoc sp. strains. All except two strains of Leuconsotoc mesenteroides subsp. mesenteroides, as well as two strains formerly identified as L. gelidum, and one strain of Leuconostoc showed a common band at about 1.1 Kb. This significant band was absent in all other strains of the genus Leuconostoc. In conclusion, in this study the set of primers from RAPD-PCR allowed identification of strains belonging to L. mesenteroides subsp. mesenteroides, and could be used for biotechnological purposes to monitor this microorganism in mixed populations of microflora present in fermented feed and food products. Similar conclusions were obtained in another study in which a reliable, reproducible, and rapid molecular RAPDPCR fingerprinting method was developed to identify the three subspecies of Leuconostoc mesenteroides (mesenteroides, cremoris, and dextranicum). rRNA-Based Techniques. Ribosomal RNA-based techniques including analysis of profiles generated by ISR amplification, ISR restriction, and ARDRA were evaluated to identify Leuconostoc. In a study by Chenoll et al.,71 13 Leuconostoc isolates were included and challenged for characterization by rRNA-based techniques. These include L. argentinum, L. carnosum, L. citreum, L. fallax, L. ficulneum, L. fructosum, L. gasicomitatum, L. gelidum, L. lactis, L. mesenteroides supsp. cremoris, L. mesenteroides sups. dextranicum, L. mesenteroides subsp. mesenteroides, and L. pseudomesenteroides. The 16S-23S ISR amplified from genomic DNA in all Leuconostoc strains, except L. fallax, shared a single ISR amplified fragment of about 700 bp, which showed an ISR amplification pattern with four bands of sizes 1450, 830, 730, and 650 bp. ISR restriction analysis with DdeI of Leuconostoc revealed four to six bands. Six species with 93% similarity showed single profiles, whereas the remainder were clustered in two groups. One group included L. gelidum and L. gasicomitatum, and the other group clustered L. lactis and L. citreum together with the three subspecies of L. mesenteroides. The Hae III restriction enzyme failed to digest the amplified ISR of L. fallax, whereas ISR-Hae III band patterns of the remaining Leuconostoc species yielded between one and five bands. Seven profiles with 90% similarity were included in a single cluster and the other grouped into two clusters. One cluster included L. gelidum and L. pseudomesenteroides.

Molecular Detection of Human Bacterial Pathogens

The three subspecies of L. mesenteroides shared a profile with L. gasicomitatum. On the other hand, the amplified 16S ribosomal DNA restriction analysis (ARDRA) with Hae III of the abovementioned strains of Leuconostoc generated seven profiles of two to four bands. Some of these band patterns were shared by a number of strains: (i) L. ficulneum and L. fructosum; (ii) L. carnosum, L. gelidum, and L. lactis; (iii) L. pseudomesenteroides and its three subspecies. Leuconostoc argentinum, L. citreum, L. fallax, and L. gasicomitatum also yielded specific band patterns. When ARDRA was performed with Dde I, four unique profiles were recognized (L. argentinum, L. carnosum, L. fallax, and L. fructosum), along with a profile that was shared by L. ficulneum, L. gelidum, and L. mesenteroides subsp. mesenteroides, and another profile that was the same for the rest of strains. When the method was used with both restriction enzymes (HaeIII and DdeI) all species were discriminated, except L. pseudomesenteroides, which was obtained by the BioNumerics Composite Data Set tool. Epidemiological Studies. Leuconostoc spp. are rarely isolated from hospital environments because of their epidemiology and virulence features. Very few reports of hospitalassociated cases associated with Leuconostoc spp. have been described in the relevant literature.50,72,73 In the first example, L. pseudomesenteroides was described as a cause of nosocomial urinary tract infections in five patients.73 Analysis of chromosomal DNA by pulsedfield-gel electrophoresis (PFGE) after treatment with SmaI indicated a clonal relationship among the isolates, and the report emphasized the possibility of nosocomial transmission of this unusual opportunistic, vancomycin-resistant pathogen. In another example, 42 patients became infected between July 2003 and October 2004 by strains of L. mesenteroides subsp. mesenteroides (genotype 1) in different departments of the Juan Canalejo Hospital in northwest Spain. During 2006, six inpatients in different departments of the same hospital became infected (genotypes 2–4) by the same L. mesenteroides subsp. mesenteroides. After an epidemiological study and molecular analysis by PFGE with ApaI as restriction enzyme, it was concluded that parenteral nutrition was the most likely source of infection, and it was suggested that products manufactured in the centralized hospital pharmacy may have been the origin of the unexpected nosocomial outbreak.50

25.2   METHODS 25.2.1  Sample Preparation Sample Collection and Culture. Freshly collected fecal samples are diluted tenfold in phosphate buffer (0.05 M pH 7.0) and stored at −82°C in 1-ml aliquots for later DNA extraction. For bacteriological culture, fresh fecal samples are homogenized and diluted tenfold in prereduced diluent (containing 8.5 g of NaCl, 1.0 g of peptone, and 0.1 g of cysteine per liter, pH 7.0) in an anaerobic chamber. Viable cell counts

275

Leuconostoc

of lactobacilli are determined by plating on Rogosa SL agar (Difco) after incubation for 2 days. Colonies are picked randomly from an agar plate containing 30–300 colonies. These subcultured isolates represent the predominant strains comprising the population selected on Rogosa agar.74 DNA Extraction. Frozen fecal sample (1 mL) is thawed on ice and centrifuged. The pellet is resuspended in 100 μL of lysis buffer (6.7% sucrose, 50 mM Tris-HCl pH 8.0, 10 mM EDTA, 20 mg of lysozyme per mL, 1000 U of mutanolysin per ml, 100 μg of RNaseA per ml). After incubation for 1 h at 37°C, 6 μL of sodium dodecyl sulfate (20%) and 5 μL of proteinase K solution (15 mg/mL) are added, and the mixture was further incubated for 15 min at 60°C until the cells lysed. After cooling on ice, 400 μL of Tris-HCl (pH 8.0) is added and the mixture is extracted once with phenol-chloroformisoamyl alcohol (25:24:1) and twice with chloroform. After ethanol precipitation the DNA is dissolved in 100 μL of Tris HCl pH 8.0).74 To extract DNA from cultured isolates, 2 mL of inoculated culture medium is incubated overnight at 30°C, and cells are concentrated by centrifugation at 4500 × g for 5  min. The pellet is resuspended in 180 μL lysis buffer (20 mM TrisHCl, pH 8.0, 2 mM EDTA, 1.2% Triton X-100, 20 mg/mL lysozyme, 100 U/mL mutanolysin) and incubated for 30 min at 37°C. Afterwards, the DNeasy tissue kit (QIAGEN) is applied.73





25.3  CONCLUSION Although Leuconostoc spp. are present in dairy products and are relatively nonpathogenic to human hosts, infections with these bacteria do occur occasionally, especially in immunocompromised individuals, pediatric patients, and the elderly who have a catheter or ventilator implanted. The clinical symptoms range from bacteremia/sepsis, brain and liver abscesses, peritonitis, endophthalmitis, endocarditis, osteomylitis, to nosocomial urinary tract infections. Recent development and application of molecular diagnostic procedures have done much to give Leuconostoc spp. long-overdue recognition as important opportunistic pathogens. It is envisaged that with their high sensitivity, specificity, and rapid result availability, molecular tests will play an even more prominent role in the diagnosis and management of clinical diseases due to Leuconostoc spp. in future.

REFERENCES

25.2.2  Detection Procedures We present below a real-time PCR protocol for detection and quantitation of Leuconostoc spp. that was developed by Friedrich and Lenke (2006).73 Other PCR procedures may be also utilized.68,74–80 The quantitative real-time PCR of Friedrich and Lenke73 are based on 16S rRNA gene primers and probe.



Primer or Probe

Sequence (5′→3′)

Bac1108R

CGCTCGTTGCGGGAC

Leuc986F

CAGGTCTTGACATCCTTTGAAG

PLeuc1026F

Cy5-CTCTTCGGAGACAAAGTGA CAGGT-BHQ2

Target Bacterial 16S rRNA gene Leuconostoc spp. 16S rRNA gene Leuconostoc spp. 16S rRNA gene







Procedure 1. Oligonucleotides are diluted in ultraPure distilled water (Invitrogen) to stock concentrations of 100 μM. 2. PCR mixture (20 μL) is composed of 1× PCR buffer (QIAGEN), 5 mM MgCl2, 0.9 μM of each of the primers (Leuc986F and Bac1108R), 0.2 μM of the dual-labeled probe PLeuc1026F, 0.8 mM of each dNTPs, 2.5 U of HotStarTaq DNA polymerase (QIAGEN), and 4 μL of template.

3. Amplification and detection are carried out on a Rotor-Gene model 3000 quantitative PCR cycler (Corbett Life Science), applying a first denaturation at 95°C for 15 min, followed by 50 cycles at 94°C for 20 s and 58°C for 45 s. 4. Fluorescence data are acquired at the end of each elongation step at 58°C in the channel Cy5.





1. Garvie, E.I., Genus Leuconostoc. In Sneath, P.H.A., Mair, N.S., and Sharp, M.E. (Editors), Bergey’s Manual of Systematic Bacteriology, Vol. 2, pp. 1075–1079, The Williams & Wilkins Co., Baltimore, 1986. 2. Benkerroum, N. et al., Development and use of a selective medium for isolation of Leuconostoc spp. from vegetables and dairy products, Appl. Environ. Microbiol., 59, 607, 1993. 3. Elliott, J.A., and Facklam, R.R., Identification of Leuconostoc spp. by analysis of soluble whole-cell protein patterns, J. Clin. Microbiol., 31, 1030, 1993. 4. Teuber, M., The family Streptoccaceae (nonmedical aspects). In Starr, M.P. (Editor), The Prokaryotes, Vol. 2, pp. 1614– 1630, Springer-Verlag, New York, 1981. 5. Facklam, R., and Elliott, J.A., Identification, classification, and clinical relevance of catalase-negative, gram-positive cocci, excluding the streptococci and enterococci, Clin. Microbiol. Rev., 8, 479, 1995. 6. Ruoff, K.L., Aerococcus, Abiotrophia, and other aerobic catalase-negative, gram-positive cocci. In Murray, P.R., Baron, E.J., Jorgensen, J.H., Landry, M.L., and Pfaller, M.A. (Editors), Manual of Clinical Microbiology, Vol. 1, 31: 443– 454, ASM PRESS, Washington, DC, 2007. 7. Christensen, J.J., and Facklam, R.R., Granulicatella and Abiotrophia species from human clinical specimens, J. Clin. Microbiol., 39, 3520, 2001. 8. Collins, M.D., and Lawson, P.A., The genus Abiotrophia (Kawamura et al.) is not monophyletic: Proposal of Granulicatella gen. nov., Granulicatella adiacens comb. nov., Granulicatella elegans comb. nov. and Granulicatella balaenopterae comb. nov., Int. J. Syst. Evol. Microbiol., 50, 365, 2000. 9. Kawamura, Y. et al., Transfer of Streptococcus adjacens and Streptococcus defectivus to Abiotrophia gen. nov. as Abiotrophia adiacens comb. nov. and Abiotrophia defectiva comb. nov., respectively, Int. J. Syst. Bacteriol., 45, 798, 1995.

276 10. Collins, M.D. et al., Aerococcus christensenii sp. nov., from the human vagina, Int. J. Syst. Bacteriol., 49, 1125, 1999. 11. Facklam, R. et al., Phenotypic description and antimicrobial susceptibilities of Aerococcus sanguinicola isolates from human clinical samples, J. Clin. Microbiol., 41, 2587, 2003. 12. Lawson, P.A. et al., Aerococcus sanguicola sp. nov., isolated from a human clinical source, Int. J. Syst. Evol. Microbiol., 51, 475, 2001. 13. Christensen, J.J. et al., Aerococcus-like organism, a newly recognized potential urinary tract pathogen, J. Clin. Microbiol., 29, 1049, 1991. 14. Christensen, J.J. et al., Aerococcus urinae: intraspecies genetic and phenotypic relatedness, Int. J. Syst. Bacteriol., 47, 28, 1997. 15. Lawson, P.A. et al., Aerococcus urinaehominis sp. nov., isolated from human urine, Int. J. Syst. Evol. Microbiol., 51, 683, 2001. 16. Collins, M.D. et al., Dolosicoccus paucivorans gen. nov., sp. nov., isolated from human blood, Int. J. Syst. Bacteriol., 49, 1439, 1999. 17. Aguirre, M., and Collins, M.D., Phylogenetic analysis of Alloiococcus otitis gen. nov., sp. nov., an organism from human middle ear fluid, Int. J. Syst. Bacteriol., 42, 79–83, 1992. 18. LaClaire, L.L., and Facklam, R.R., Comparison of three commercial rapid identification systems for the unusual gram-positive cocci Dolosigranulum pigrum, Ignavigranum ruoffiae, and Facklamia species, J. Clin. Microbiol., 38, 2037, 2000. 19. Lawson, P.A. et al., Facklamia languida sp. nov., isolated from human clinical specimens, J. Clin. Microbiol., 37, 1161, 1999. 20. Collins, M.D. et al., Phenotypic and phylogenetic characterization of some Globicatella-like organisms from human sources: Description of Facklamia hominis gen. nov., sp. nov., Int. J. Syst. Bacteriol., 47, 880, 1997. 21. Collins, M.D. et al., Facklamia sourekii sp. nov., isolated from human sources, Int. J. Syst. Bacteriol., 49, 635, 1999. 22. Collins, M.D. et al., Facklamia ignava sp. nov., isolated from human clinical specimens, J. Clin. Microbiol., 36, 2146, 1998. 23. Collins, M.D. et al., Gemella bergeriae sp. nov., isolated from human clinical specimens, J. Clin. Microbiol., 36, 1290, 1998. 24. Collins, M.D. et al., Description of Gemella sanguinis sp. nov., isolated from human clinical specimens, J. Clin. Microbiol., 36, 3090, 1998. 25. Collins, M.D. et al., Globicatella sanguis gen.nov., sp.nov., a new gram-positive catalase-negative bacterium from human sources, J. Appl. Bacteriol., 73, 433, 1992. 26. Facklam, R., What happened to the streptococci: Overview of taxonomic and nomenclature changes, Clin. Microbiol. Rev., 15, 613, 2002. 27. Shewmaker, P.L. et al., DNA relatedness, phenotypic characteristics, and antimicrobial susceptibilities of Globicatella sanguinis strains, J. Clin. Microbiol., 39, 4052, 2001. 28. Collins, M.D. et al., Phylogenetic analysis of some Aerococcus-like organisms from clinical sources: Description of Helcococcus kunzii gen. nov., sp. nov., Int. J. Syst. Bacteriol., 43, 425, 1993. 29. Collins, M.D. et al., Ignavigranum ruoffiae sp. nov., isolated from human clinical specimens, Int. J. Syst. Bacteriol., 49, 97, 1999. 30. Flaherty, J.D. et al., Fatal case of endocarditis due to Weissella confusa, J. Clin. Microbiol., 41, 2237, 2003.

Molecular Detection of Human Bacterial Pathogens 31. Olano, A. et al., Weissella confusa (basonym: Lactobacillus confusus) bacteremia: A case report, J. Clin. Microbiol., 39, 1604, 2001. 32. Collins, M.D. et al., Helcococcus sueciensis sp. nov., isolated from a human wound, Int. J. Syst. Evol. Microbiol., 54, 1557, 2004. 33. Panackal, A.A. et al., Prosthetic joint infection due to “Helcococcus pyogenes” [corrected], J. Clin. Microbiol., 42, 2872, 2004. 34. Ruoff, K.L. et al., Vancomycin-resistant gram-positive bacteria isolated from human sources, J. Clin. Microbiol., 26, 2064, 1988. 35. Garvie, E.I., Leuconostoc oenos sp.nov., J. Gen. Microbiol., 48, 431, 1967. 36. Geis, A., Singh, J., and Teuber, M., Potential of lactic streptococci to produce bacteriocin, Appl. Environ. Microbiol., 45, 205, 1983. 37. Cogan, T.M., O’Dowd, M., and Mellerick, D., Effects of pH and sugar on acetoin production from citrate by Leuconostoc lactis, Appl. Environ. Microbiol., 41, 1–8, 1981. 38. Harvey, R.J., and Collins, E.B., Role of citritase in acetoin formation by Streptococcus diacetilactis and Leuconostoc citrovorum, J. Bacteriol., 82, 954, 1961. 39. Edwards, C.G. et al., Lactobacillus kunkeei sp. nov.: A spoilage organism associated with grape juice fermentations, J. Appl. Microbiol., 84, 698, 1998. 40. Sesma, F. et al., Cloning of the citrate permease gene of Lactococcus lactis subsp. lactis biovar diacetylactis and expression in Escherichia coli, Appl. Environ. Microbiol., 56, 2099, 1990. 41. Martinez-Murcia, A.J., and Collins, M.D., A phylogenetic analysis of the genus Leuconostoc based on reverse transcriptase sequencing of 16 S rRNA, FEMS Microbiol. Lett., 58, 73, 1990. 42. Martinez-Murcia, A.J., Harland, N.M., and Collins, M.D., Phylogenetic analysis of some leuconostocs and related organisms as determined from large-subunit rRNA gene sequences: Assessment of congruence of small- and large-subunit rRNA derived trees, J. Appl. Bacteriol., 74, 532, 1993. 43. Martinez-Murcia, A.J., and Collins, M.D., A phylogenetic analysis of an atypical leuconostoc: Description of Leuconostoc fallax sp. nov., FEMS Microbiol. Lett., 66, 55, 1991. 44. Schleifer, K.H., Gram-positive cocci. In P.H.A. Sneath, NSM, M.E. Sharp, and J.G. Holt (Editors), Bergey’s Manual of Systematic Bacteriology, Vol. 2, pp. 999–1002, The Williams & Williams Co., Baltimore, 1986. 45. Kang, M.S. et al., Effect of Leuconostoc spp. on the formation of Streptococcus mutans biofilm, J. Microbiol., 45, 291, 2007. 46. Balcázar, J.L. et al., Effect of Lactococcus lactis CLFP 100 and Leuconostoc mesenteroides CLFP 196 on Aeromonas salmonicida Infection in brown trout (Salmo trutta), J. Mol. Microbiol. Biotechnol., 17, 153, 2009. 47. Handwerger, S. et al., Infection due to Leuconostoc species: Six cases and review, Rev. Infect. Dis., 12, 602, 1990. 48. Ling, M.L., Leuconostoc bacteraemia, Singapore Med. J., 33, 241, 1992. 49. Camarasa, A., Chiner, E., and Sancho-Chust, J.N., Pulmonary abscess due to Leuconostoc species in an immunocompetent patient, Arch. Bronconeumol., 45, 471, 2009. 50. Bou, G. et al., Nosocomial outbreaks caused by Leuconostoc mesenteroides subsp. mesenteroides, Emerg. Infect. Dis., 14, 968, 2008.

Leuconostoc 51. Dhodapkar, K.M., and Henry, N.K., Leuconostoc bacteremia in an infant with short-gut syndrome: Case report and literature review, Mayo Clin. Proc., 71, 1171, 1996. 52. Florescu, D. et al., Leuconostoc bacteremia in pediatric patients with short bowel syndrome: Case series and review, Pediatr. Infect. Dis. J., 27, 1013, 2008. 53. Jofre, M.L. et al., Leuconostoc infections in patients with short gut syndrome, parenteral nutrition and continuous enteral feeding, Rev. Chil. Infectol., 23, 340, 2006. 54. Bernaldo de Quiros, J.C. et al., Leuconostoc species as a cause of bacteremia: Two case reports and a literature review, Eur. J. Clin. Microbiol. Infect. Dis., 10, 505, 1991. 55. Isenberg, H.D. et al., Clinical laboratory challenges in the recognition of Leuconostoc spp., J. Clin. Microbiol., 26, 479, 1988. 56. Rubin, L.G. et al., Infection with vancomycin-resistant “streptococci” due to Leuconostoc species, J. Infect. Dis., 157, 216, 1988. 57. Peters, V.B. et al., Leuconostoc species bacteremia in a child with acquired immunodeficiency syndrome, Clin. Pediatr. (Phila), 31, 699, 1992. 58. Coovadia, Y.M., Solwa, Z., and van den Ende, J., Meningitis caused by vancomycin-resistant Leuconostoc sp., J. Clin. Microbiol., 25, 1784, 1987. 59. Friedland, I.R., Snipelisky, M., and Khoosal, M., Meningitis in a neonate caused by Leuconostoc sp., J. Clin. Microbiol., 28, 2125, 1990. 60. Montejo, M. et al., Abdominal abscess due to Leuconostoc species in a liver transplant recipient, J. Infect., 41, 197, 2000. 61. Zaoui, A. et al., Leuconostoc osteomyelitis, Joint Bone Spine, 72, 79, 2005. 62. Deye, G. et al., A case of Leuconostoc ventriculitis with resistance to carbapenem antibiotics, Clin. Infect. Dis., 37, 869, 2003. 63. Kumudhan, D., and Mars, S., Leuconostoc mesenteroids as a cause of post-operative endophthalmitis—a case report, Eye, 18, 1023, 2004. 64. Taneja, N. et al., Nosocomial urinary tract infection due to Leuconostoc mesenteroides at a tertiary care centre in north India, Indian J. Med. Res., 122, 178, 2005. 65. Swenson, J.M., Facklam, R.R., and Thornsberry, C., Anti­ microbial susceptibility of vancomycin-resistant Leuco­ nostoc, Pediococcus, and Lactobacillus species, Antimicrob. Agents Chemother., 34, 543, 1990. 66. Golan, Y. et al., Daptomycin for line-related Leuconostoc bacteraemia, J. Antimicrob. Chemother., 47, 364, 2001. 67. de la Maza, L., Ruoff, K.L., and Ferraro, M.J., In vitro activities of daptomycin and other antimicrobial agents against vancomycin-resistant gram-positive bacteria, Antimicrob. Agents Chemother., 33, 1383, 1989.

277 68. Lee, H.J., Park, S.Y., and Kim, J., Multiplex PCR-based detection and identification of Leuconostoc species, FEMS Microbiol. Lett., 193, 243, 2000. 69. Cusick, S.M., and O´Sullivan, D., Use of single, triplicate arbitrarily primed-PCR procedure for molecular fingerprinting of lactic acid bacteria, Appl. Environ. Microbiol., 66, 2227, 2000. 70. Moschetti, G. et al., Specific detection of Leuconostoc mesenteroides subs. mesenteroides with DNA primers identified by randomly amplified polymorphic DNA analysis, Appl. Environm. Microbiol., 66, 422, 2000. 71. Chenoll, E., Macian, M.C., and Aznar, R., Identification of Carnobacterium, Lactobacillus, Leuconostoc and Pediococcus by rDNA-based tecniques, Syst. Appl. Microbiol., 26, 546, 2003. 72. Scano, F. et al., Leuconostoc species: A case-cluster hospital infection, Scand. J. Infect. Dis., 31, 371, 1999. 73. Cappelli, E.A. et al., Leuconostoc pseudomesenteroides as a cause of nosocomial urinary tract infections, J. Clin. Microbiol., 37, 4124, 1999. 74. Friedrich, U., and Lenke, J., Improved enumeration of lactic acid bacteria in mesophilic dairy starter cultures by using multiplex quantitative real-time PCR and flow cytometryfluorescence in situ hybridization, Appl. Environ. Microbiol., 72, 4163, 2006. 75. Walter, J. et al., Detection of Lactobacillus, Pediococcus, Leuconostoc, and Weissella species in human feces by using group-specific PCR primers and denaturing gradient gel electrophoresis, Appl. Environ. Microbiol., 67, 2578, 2001. 76. Heilig, H.G. et al., Molecular diversity of Lactobacillus spp. and other lactic acid bacteria in the human intestine as determined by specific amplification of 16S ribosomal DNA, Appl. Environ. Microbiol., 68, 114, 2002. 77. Jang, J. et al., A rapid method for identification of typical Leuconostoc species by 16S rDNA PCR-RFLP analysis, J. Microbiol. Methods, 55, 295, 2003. 78. Walczak, P., Konopacka, M., and Otlewska, A., Genetic diversity among Lactococcus sp. and Leuconostoc sp. strains using PCR-RFLP of insertion sequences ISS1-type, IS904, and IS982, Pol. J. Microbiol., 54, 183, 2005. 79. Elizaquível, P., Chenoll, E., and Aznar, R., A TaqMan-based real-time PCR assay for the specific detection and quantification of Leuconostoc mesenteroides in meat products, FEMS Microbiol. Lett., 278, 62, 2008. 80. Schillinger, U. et al., A genus-specific PCR method for differentiation between Leuconostoc and Weissella and its application in identification of heterofermentative lactic acid bacteria from coffee fermentation, FEMS Microbiol. Lett., 286, 222, 2008.

26 Listeria Dongyou Liu and Ting Zhang CONTENTS 26.1 Introduction...................................................................................................................................................................... 279 26.1.1 Classification, Morphology, and Genome Organization...................................................................................... 279 26.1.2 Biology and Epidemiology................................................................................................................................... 282 26.1.3 Clinical Features and Pathogenesis...................................................................................................................... 283 26.1.4 Diagnosis.............................................................................................................................................................. 284 26.1.4.1 Phenotypic Techniques.......................................................................................................................... 284 26.1.4.2 Genotypic Techniques............................................................................................................................ 286 26.2 Methods............................................................................................................................................................................ 287 26.2.1 Sample Preparation............................................................................................................................................... 287 26.2.2 Detection Procedures............................................................................................................................................ 287 26.2.2.1 Multiplex PCR Identification of Listeria Species.................................................................................. 287 26.2.2.2 Multiplex PCR Differentiation of L. monocytogenes Serogroups......................................................... 288 26.2.2.3 Multiplex PCR Determination of L. monocytogenes Virulence........................................................... 288 26.2.2.4 Multiplex PCR Determination of L. monocytogenes Serotypes 1/2a and 4b and Epidemic Clones I, II, and III.............................................................................................................................. 289 26.2.2.5 Real-Time PCR Detection of L. monocytogenes................................................................................... 290 26.3 Conclusion and Future Perspectives................................................................................................................................. 290 References.................................................................................................................................................................................. 291

26.1  INTRODUCTION 26.1.1  Classification, Morphology, and Genome Organization Classification. The genus Listeria is classified taxonomically in the family Listeriaceae, order Bacillales, class Bacilli, phylum Furmicutes. The only other genus in the family Listeriaceae is Brochothrix (consisting of two species, Brochothrix campestris and Brochothrix thermosphacta). Until recently, six closely related, gram-positive bacterial species have been recognized in the genus Listeria—that is, L. monocytogenes, L. ivanovii, L. seeligeri, L. innocua, L. welshimeri, and L. grayi, with L. ivanovii being further separated into two subspecies, L. ivanovii subsp. londoniensis and L. ivanovii subsp. ivanovii.1,2 Whereas L. monocytogenes is a facultative intracellular pathogen for humans and animals, L. ivanovii (formerly L. monocytogenes serotype 5) mainly infects ungulated animals (e.g., sheep and cattle), and the other four species are essentially saprophytes that have adapted for free living in soil and decaying vegetation.1,3 In late 2009, two additional Listeria species were described. Having originated from soil and water samples at the Finger Lakes National Forest in New York, L. marthii sp. nov. demonstrates a close phylogenetic relatedness to L. monocytogenes and L. innocua, and a more distant relatedness to

L. welshimeri, L. seeligeri, L. ivanovii, and L. grayi. Given its absence of a homologue of the L. monocytogenes virulence gene island, L. marthii is likely nonpathogenic.4 L. rocourtiae sp. nov. was identified from precut lettuce in Austria. This species is distinguishable from other Listeria species by using phenotypic tests, and its type strain is avirulent as assessed by in vitro cell culture and in vivo mouse model.5 On the basis of serological interactions between Listeria somatic (O)/flagellar (H) antigens and their corresponding antisera, Listeria is differentiated 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; L. welshimeri of serovars 6a and 6b; L. seeligeri of serovars 1/2b, 4c, 4d, and 6b; and L. grayi of serovar Grayi (Table 26.1).6 Upon phylogeneitc analyses, L. monocytogenes is grouped into three evolutionary 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.7,8 It was recently shown that L. monocytogenes lineage III can be subdivided into three distinct genetic subgroups, IIIA, IIIB, and IIIC, with subgroup IIIA consisting of typical rhamnose-positive avirulent serovar 4a and virulent serovar 4c strains; subgroup IIIC of atypical rhamnosenegative virulent serovar 4c strains; and subgroup IIIB of 279

280

Molecular Detection of Human Bacterial Pathogens

of human listeriosis, with serovar 4b alone being responsible for 49% of Listeria-related foodborne diseases during a survey of French patients between 2001 and 2003 (Table 26.3).16 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. However, all L. monocytogenes serovars except 4a have the capability to cause mouse mortality via intraperitoneal route (Table 26.4).9,10,12,17,18 Morphology. Listeria is a gram-positive, nonsporeforming, motile, facultatively anaerobic, rod-shaped bacterium of 0.4–0.5 µm × 1–1.5 µm. In direct smears, Listeria may appear coccoid, resembling streptococci, and longer cells may look like corynebacteria. This bacterium exhibits characteristic tumbling motility when examined by light microscopy. The organism is actively motile by means of peritrichous flagella at room temperature (20°C–25°C), but it does not synthesize flagella at body temperatures (37°C). Being nutritionally undemanding, Listeria spp. grow well on a number of nonselective microbiological media such as tryptone soy broth (TSB) 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 their optimal growth. Colonies of 0.5–1.5 mm in diameter usually emerge after 24–48 h incubation on agar media (e.g., blood agar or BHI agar), and appear as round, translucent, low convex with a smooth surface and a crystalline center. After 3–7 days, older and larger colonies of 3–5 mm in diameter may give 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

TABLE 26.1 Compositions of Somatic (O)/Flagellar (H) Antigens in Listeria Serovars Serovar

O-antigen

1/2a 1/2b 1/2c 3a 3b 3c 4a 4ab 4b 4c 4d 4e 7 5 6a 6b

H-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

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

atypical rhamnose-negative virulent nonserovar 4a and nonserovar 4c strains, some of which may be related to serovar 7 (Table 26.2).9–14 Subsequent study revealed that subgroup IIIB (including serovar 7) forms a separate lineage (IV) within L. monocytogenes.15 In accordance with its virulence potential in mammalian hosts, L. monocytogenes is also separated into pathogenic (virulent) and nonpathogenic (avirulent) strains. It is most noteworthy that L. monocytogenes serovars 4b, 1/2a, 1/2b, and 1/2c account for over 98% isolations from clinical cases

TABLE 26.2 Characteristics of L. monocytogenes Lineages I–IVa

Lineage

Serovar

I

1/2b 3b 4b 4d 4e 1/2a 1/2c 3a 3c 4a 4c 4c 7 and unusual 4a, 4b, 4c

II

IIIA IIIC IV (IIIB)

a b

PCR Reactivity

Rhamnose Activity

inlA

lmo0733

lmo2672

inlJ

inlCb

lmo1134

ORF2819

ORF2110

lmo0737

lmo1118

+ + + + + + + + + + + − −

+ + + + + + + + + + + + +

+ + + + + + + + + + + + −

+ + + + + + + + + − − + +

+ + + + + + + + + − + + −

+ + + + + + + + + − ± + +

+ + + + + + + + + − − − −

+ + + + + − − − − − − − +

− − + + + − − − − − − − −

− − − − − + + + + − − − −

− − − − − − + − + − − − −

Summarized from Liu et al.9–12; Doumith et al.13; Roberts et al.14 inlC is also found in some L. ivanovii strains.12

281

Listeria

(SLCC 5334) (Table 26.5). Interestingly, the nonpathogenic L. seeligeri type strain (of serovar 1/2b) (SLCC3954) harbors the smallest genome of the Listeria species sequenced to date.22 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 strainspecific genes are identifiable, respectively, from L. monocytogenes serovar 1/2a strains EGD-e and F6854, and from serovar 4b strains F2365 and H7858 (Table 26.5). 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 species-specific genes are recognized in L. innocua CLIP 11262 and L. welshimeri SLCC 5334, respectively (Table 26.5).19–21 The fact that some L. monocytogenes strain-specific genes tend to have atypical base composition implies their acquisition through horizontal gene transfer. In addition, some L.  monocytogenes strain-specific genes encode putative surface-associated proteins such as internalins, which may contribute to the increased pathogenicity of the strains concerned.20 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.20 While the genomes of serovar 4b strains (F2365 and H7858) do not contain intact insertion sequence (IS)

TABLE 26.3 L. monocytogenes Serovars Causing Human Listeriosisa Serotype

No. of Isolates (%)

4b

294/603 (49%)

1/2a

163/603 (27%)

1/2b

120/603 (20%)

1/2c

22/603 (4%)

3a/3b

4/603 ( M/N diseases > Bacteremia Bacteremia > M/N diseases > CNS infections M/N diseases > Bacteremia > CNS infections Bacteremia > CNS infections > M/N diseases Bacteremia

Based on the analysis of 603 L. monocytogenes isolates from 603 French patients during 2001–2003.16 M/N diseases, maternal-neonatal diseases; CNS infections, central nervous system infections. a

to retain the Gram stain and fail to display characteristic violet color. The optimal growth temperatures for Listeria are between 30°C and 37°C; Listeria are motile with peritrichous flagella at 20°C–28°C, but become nonmotile at 37°C. Genomic Organization. The genus Listeria possesses genomes of relatively low G + C contents of 36%–39%, which implies its relatedness to other members of the phylum Firmicutes such as the genera Bacillus, Brochothrix, Clostridium, Enterococcus, Lactobacillus, Staphylococcus, and Streptococcus. The recent publication of whole genome sequences of several Listeria spp. have helped uncover valuable new details on the differential genetic compositions among Listeria bacteria.19–22 These include two L. monocytogenes serovar 1/2a (EGD-e and F6854) and two serovar 4b (F2365 and H7858) strains, one L. innocua (CLIP 11262), one L. seeligeri (SLCC3954), and one L. welshimeri strain TABLE 26.4 L. monocytogenes Serovars and Relative Virulence

PCRa Strain HCC8 EGD ATCC 19112 ATCC 19114 HCC25 ATCC 19115 ATCC 19116 874 ATCC 19117 ATCC 19118 R2-142 a b c

Source Catfish brain Guinea pig Human Ruminant brain Catfish kidney Human Chicken Cow brain Sheep Chicken Food

Serovar 1 1/2a 2 4a 4a 4b 4c 4c 4d 4e 7

inlJ

inlC

+ + + − − + + + + + −

+ + + − − + − + + + +

LD50b

Relative Virulence (%)c

3 3

Streptococcus sp.

Anaerobic

Bacteroides uniformis Fusobacterium sp. Bacteroides fragilis group 2

Bacteroides fragilis >3 2 >3 >3 >3 2 Bacteroides distasonis >3 0 Prevotella sp. >3 >3 3 >3 0 >3 2 Bacteroides thetaiotaomicron

Bacteroides vulgatus

Data are from Reference 4 and Jean-Pierre, H., Jumas-Bilak, E., and Marchandin H., unpublished data, 2009. Strains labeled “ADV” were previously published.4 Number of species indicated in case of polymorphic aerobic and/or anaerobic cultures; species name indicated in case of pure culture. Main associated species were Bacteroides spp., mainly species of the B. fragilis group; Enterobacteriaceae, mainly E. coli, and Enterococcus spp. Species identification: >99% of 16S rRNA gene sequence identity level with type strain of the species. F, female; M, male; ICU, intensive care unit; CNS, coagulase-negative Staphylococcus.

amoxicillin + clavulanate as well as to ticarcillin + clavulanate was observed.40 Described in 2000, ACI-1 remains the only β-lactamase hitherto described among strict anaerobic gramnegative cocci. The G + C content of aci1 gene (42.1 mol%) differed from that of A. fermentans total DNA (54.7–57.4 mol%) suggesting that this gene was probably acquired by horizontal gene transfer. Highest sequence similarity levels were observed with class A β-lactamases from Haemophilus influenzae and from some gram-positive organisms, particularly

belonging to the genus Bacillus. Further investigations on flanking regions of the aci1 gene revealed the existence of transposable genetic elements in the Acidaminococcus strain (transposase and putative resolvase), suggesting that this gene may be part of a transposable element.44 In a large study on ertapenem activity on anaerobes, Goldstein et al. reported that one out of the two A. fermentans included in their work displayed resistance to ertapenem with minimum inhibitory concentration ≥16 μg/mL.38

345

Acidaminococcus

30.1.6  Diagnosis Conventional Techniques. Strictly anaerobic conditions are required for growth of the Acidaminococcus species. For example, Acidaminococcus spp. grew well in anaerobic conditions obtained in an anaerobic jar with AnaeroGen System® (Oxoid, Basingstoke, UK). No enrichment method and no specific selective medium exist for growth of Acidaminococcus spp.1,4,9,11 Many agar media or broths can be used for cultivation and isolation of Acidaminococcus spp. Rogosa described the growth of A. fermentans on Reinforced Clostridial Medium (RCM; Oxoid, Basingstoke, UK).1 Sugihara et al. isolated A. fermentans on several media including kanamycin-vancomycin blood agar, neomycin blood agar, Veillonella agar with neomycin, Fusobacterium agar, blood agar, China-blue agar, and rifampin-vancomycin blood agar.20 However, in the routine analysis of human clinical samples, Acidaminococcus spp. isolates grew well on Columbia sheep blood agar incubated at 37°C during 2–5 days. Both species yielded nonpigmented and nonhemolytic colonies on this medium, and colony morphology is as follows after 48 h of incubation at 37°C: for A. fermentans, colonies are 1–2 mm in diameter, round, entire, slightly raided, and whitish gray or nearly transparent; for A. intestini, colonies are about 0.5 mm in diameter, circular, convex, and whitish with a smooth surface.1,4 Members of the genus Acidaminococcus are gas producers. Other biochemical tests are mainly negative for these species. Acidaminococcus spp. do not possess oxidase nor catalase. Urease activity, nitrate reduction, gelatin liquefaction, milk modification, and aesculin hydrolysis are negative. H2S is not produced. Indole is generally not produced.1,4,9,11 Presumptive identification tests showed that all the strains displayed susceptibility to discs that contained 4 μg metronidazole or 10 μg colistin (Rosco, Taastrup, Denmark) and resistance to discs that contained 5 μg vancomycin. Susceptibility to discs that contained 1 mg kanamycin or 1 mg bile is noted for strains of A. fermentans and for the majority of A. intestini isolates. However, A. intestini strains with resistance to kanamycin or bile discs were described.4 Using rapid ID 32 A system, Acidaminococcus species displayed the following positive enzymatic reactions: arginine arylamidase, leucine arylamidase, glycine arylamidase, and histidine arylamidase activities. Two enzymatic reactions may differentiate the two species of the genus, pyroglutamic acid arylamidase, and leucyl glycine arylamidase activities.4 The metabolic end products produced by both species are acetic and butyric acids. In addition, A. intestini strains produced propionic acid and variable amounts of lactic acid, 2-hydroxybutyric, and 2-hydroxyvaleric acids.1,4 Several of the previously cited characteristics were found to be variable among strains of a same species making phenotypic identification difficult. In fine, phenotypic identification of anaerobic gram-negative cocci is mainly based on nitrate reduction detection, which allowed the identification of the genus Veillonella mainly encountered in human

clinical samples. Anaerobic gram-negative cocci, which did not reduce nitrate, are often submitted to molecular identification.33 Molecular Techniques. Molecular methods for identification included 16S and 23S rRNA genes sequencing that may be performed on DNA extracts rapidly obtained by a boiling–freezing method.4 The two species of the genus Acidaminococcus each formed a very tight group on the basis of their rRNA gene sequences. Indeed, intraspecific sequences identity level was >99.5% for both species and both genes. Interspecific 16S and 23S rRNA gene sequence identity levels were 99% identity to C. hongkongensis (GenBank accession no. AJ318864). However, the true identity and disease association of this isolate has not been reported. It is likely that this isolate is also a strain of C. hongkongensis that has been misidentified as a Ruminococcus strain. The rare report of C.  hongkongensis may be a reflection of the difficulty in accurately identifying anaerobic gram-positive bacilli in clinical laboratories. Now that 16S rRNA gene analysis is more readily available, more strains of C. hongkongensis will be uncovered, which will help better define its epidemio­ logy and clinical spectrum of disease. Since the first report of C. hongkongensis infections, 16S rRNA sequence related to C. hongkongensis has subsequently been detected in various environmental samples. In a study on urban aerosols collected in the United States, 16S rRNA sequences belonging to a diverse bacterial population, including genus Catabacter, were detected, although the degree of sequence similarity was not mentioned.4 Similar findings were also observed in the mangrove sediment in China.5 In another study from Japan, among microbial communities from rice-paddy-field soil used for microbial fuel cells, 16S rRNA sequences of diverse bacteria, including one with 93% identity to that of C. hongkongensis.6 However, since in none of these reports were bacteria isolated for further characterization, and since the identity of these sequences with that of C. hongkongensis was not very high or unknown, it remains

to be determined if C. hongkongensis or some closely related bacteria were present in these environments. Interestingly, in a recent study on the fecal microflora of a dugong (Dugong dugong), an aquatic herbivorous mammal in captivity at Toba Aquarium, Japan, 16S rRNA sequence clones belonging to diverse bacterial phyla, including one that possessed 100% homology with C.  hongkongensis, were identified (Figure 32.1).7 This suggests that C. hongkongensis may be one of the gut commensals in this animal, which supports the speculation that the gastrointestinal tract is a source of infection in humans. In addition, C. hongkongensis 16S rRNA has been detected in lake water and anaerobic wastewater treatment reactors.8,9

32.1.4  Pathogenicity and Therapy Given its ability to invade into bloodstream causing systemic sepsis, C. hongkongensis may possess virulence factors that are yet to be defined. One possible factor may be related to its ability to produce catalase.1 Despite being strictly anaerobic, C. hongkongensis is able to survive in the presence of oxygen for 3–4 days.1 Such aerotolerance may be related to the production of catalase, which may protect C. hongkongensis from oxidative stress.1 In B. fragilis, such aerotolerance has been described as a potential virulence factor, as clinical isolates were more aerotolerant than fecal isolates.10 C. hongkongensis is most susceptible to metronidazole with minimal inhibitory concentration (MIC) of 90% of the strains had colonies >2 mm in diameter. The colonies can vary in size and color on one plate. Some colonies are convex and whitish, while others are flat and

translucent. This might give the impression that it is a mixed culture. However, colonies always have a perfect round shape, and are usually slightly convex.7 Epidemiology. F. magna is one of the most frequently recovered anaerobes from clinical infectious material. In such infections approximately 30% of the anaerobes are GPAC of which around 30% is F. magna.8 Remarkably, F. magna can be isolated from clinical specimens in pure culture and can be surpassed by Bacteroides fragilis in this aspect only. Neut et al.9 analyzed the presence of GPAC in normal oral, fecal, and vaginal microbiota. F. magna was found in the vaginal and fecal microbiota (especially healthy adults), but not in the oral microbiota. Riggio et al.10 determined the presence of F. magna in oral samples using a specific PCR. In total, 33 subgingival plaque samples of patients with adult periodontal disease and 60 pus aspirates from patients with acute dentoalveolar abscesses were analyzed. F. magna DNA was encountered in two subgingival plaque samples. These results show that F. magna is not a major pathogen in adult periodontal disease and dentoalveolar abscesses. Gao et al.11 analyzed the superficial skin bacterial biota of human forearm of six subjects. A total of 91 genera were found, of which six were observed in all subjects. One of these was Finegoldia AB109769, suggesting that F. magna is part of the normal skin microbiota. Higaki et al.12 analyzed the anaerobes isolated from infectious skin diseases. The most commonly isolated anaerobes were GPAC (66 of 106 strains, 62%), among these F. magna was the most frequently isolated GPAC (27 strains, 41%). F. magna shows pathogenic features similar to Staphylococcus aureus. In patients with infected breast abscesses, F. magna can be isolated alone or together with S. aureus.12 Since both organisms display synergism of pathogenicity, it is more difficult to cure such infections. 405

406

Molecular Detection of Human Bacterial Pathogens

Brook13 evaluated the recovery of anaerobes, among them F. magna, from clinical specimens during a 12-year period. Of all GPAC isolated, 18.4% were identified as F. magna. The majority of these F. magna strains (65.7%) were isolated from abscesses, obstetrical and gynecological infections, and wounds. The highest frequency of recovery of F. magna was shown in bone and chest infections. In pediatric patients 680 GPAC were recovered from 598 clinical specimens, from 554 patients.14 From all these strains, 10.9% were F. magna. The majority was isolated from abscesses. Bourgault et al.15 evaluated the clinical significance of F. magna in 222 patients. Of these patients, 183 had an infection in which F. magna played an important role. In 17.5% of these cases, F. magna was isolated as a pure culture from infections of bone and joint (56.3%), soft tissue (37.5%), and vascular (6.3%). In mixed infections in which F. magna involved, the most frequently isolated facultative bacteria were group D Streptococcus, Staphylococcus epidermis, Escherichia coli, and S. aureus. The most frequently found anaerobic bacteria were: Prevotella melaninogenica, B. fragilis, and Bacteroides sp. These mixed infections were mainly infections of soft tissue (37.7%), bone and joint (21.2), and foot ulcers (19.2%). From this data it can be concluded that there is a strong association between F. magna found in pure culture and orthopedic procedures and postoperative wound infection.

36.1.2  Clinical Features and Pathogenesis F. magna is capable of producing several virulence factors (Table 36.1). In 1984, Brook et al.23 examined the pathogenicity of GPAC in mixed infections. Abscesses caused by two organisms, including one strain of GPAC, were larger compared with abscesses caused by one organism. From their experiments it was concluded that F. magna and Peptostreptococcus anaerobius were equally important or more important than the other bacteria in mixed infections. This supports the hypothesis that bacteria in mixed infections may have a synergistic nature. The collagenase production of F. magna is associated with the site of infection.16

Collagen is abundantly present in the skin, tendons, and cartilage and is an organic component of bones, teeth, and the cornea. The breakdown of collagen will result in loss of tissue integrity and disease progression, thereby providing an environment suitable for growth of anaerobic bacteria. The production of collagenase may also be important for the growth of asaccharolytic bacteria, such as F. magna, since during collagen breakdown amino acids are released which may be necessary for growth and survival.24 Ng et al.25 determined the aminopeptidase activities of some GPAC strains. F. magna together with P. micra were found to degrade most substrates. There was a correlation between gelatin hydrolysis and the number of aminopeptidases produced. The authors state that gelatin hydrolysis reflects the pathogenetic potential of a strain. The growth of the bacteria can be correlated to the amount of aminopeptidases produced by protein degradation. Myhre17 described that 42% of the F. magna strains is able to bind human serum albumin (HSA). This ability was originally described for different streptococcal species. In group C and G streptococci this is mediated by protein G, and in group A streptococci by protein M.26 De Chateau et al.27 demonstrated that some strains of F. magna express protein peptostreptococcal albumin binding (PAB) on their cell surface. Sequence analyses revealed homology with the HSA-binding domain of protein G and to the framework regions of protein L (described later). This suggests an interspecies exchange of an HSA-binding protein module. In general, host binding cell wall proteins of gram-positive bacteria share a common structure, including a (from the distal NH2 terminus)28: • Signal sequence • Variable NH2-terminal region • Varying number of repeated domains that independently bind different plasma proteins • Proline-rich region supposedly intercalating the protein in the gram-positive cell wall • COOH-terminal cell wall sorting signal, required to anchor the protein to the cell wall

TABLE 36.1 Virulence Factors of F. magna Virulence Factor Collagenase PAB Protein L SufA

FAF

Function

Reference

Breakdown of collagen Binding to human serum albumin Immunoglobulin (Ig)-binding protein Release of de novo-synthesized mediators Degradation of fibrinogen Degradation of antibacterial peptides LL-37 and MIG/CXCL9 Release of FAF from bacterial cell wall Mediation of bacterial aggregation through protein-protein interactions between FAF molecules on neighboring F. magna bacteria Binding with BM40, a noncollagenous glycoprotein, present on the skin Blocking the activity of LL-37, an antibacterial peptide Inactivation of the antibacterial peptide MIG/CXCL9

16 17 18 19 20 20,21 21 22

21

Finegoldia

PAB contains a GA module (protein G-related albumin-binding module). This is a centrally located domain of 45 amino acid residues, which is responsible for the binding of HSA. This domain is subject to module shifting. The predecessor of the PAB protein is urPAB. This protein does not contain the shuffled GA module, but has a uGA domain in the NH2-terminal region. This domain shows 38% similarity with the GA module and binds HSA to a lesser extent. PAB also contains an analogous uGA domain, which indicates a second binding site for HSA. The affinity for HSA differs between the GA modules. A reason for bacteria to acquire the GA module is that the older uGA domain has lost its function due to the difference in affinity for HSA. The binding affinity for HSA is not only found on the cell surface, but also in the culture supernatant. The growth of HSA-binding strains is stimulated by the addition of HSA to the growth medium.29 This selective advantage increases the virulence of HSA-binding F. magna strains. Felten et al.30 studied the binding of 14 F. magna strains isolated from bone and joint infections to collagen, fibrinogen, and fibronectin after implantation of a foreign body. From these strains, 81% bound to collagen, 69% to fibrinogen, and 46% to fibronectin. When these results were compared to the binding abilities from F. magna strains from other infections, a correlation was found between fibrinogen binding and bone and joint infections (69% against 20%). Krepel et al.31 tried to elucidate the role of F. magna in three different polymicrobial environments: intraabdominal infections, nonpuerperal breast abscesses, and diabetic foot ulcers. An association was made between phenotypic characteristics and the site of infection. In total, 336 clinical specimens were examined: 222 were intraabdominal, from which 11 F. magna strains were isolated; 58 nonpuerperal breast abscesses, from which 21 F. magna strains were isolated; and 56 diabetic foot ulcers, from which 18 F. magna strains were isolated. From the F. magna strains the hippurate hydrolase, collagenase, and gelatinase production was determined. Strains with the lowest enzymatic activity were isolated from intraabdominal infections. The most proteolytic strains were predominantly isolated from soft-tissue infections. These are the kind of infections that tend to be chronic and heal slowly. Edmiston et al.32 showed that F. magna is the most common anaerobe isolated from nonpuerperal breast infections. F. magna strains isolated from nonpuerperal breast abscesses and diabetic foot infections were shown to have a higher collagenase production compared to F. magna strains isolated from intraabdominal infections.16 Stephens et al.33 determined the impact of the presence of GPAC present in deep tissues of chronic wounds. Clinical samples of 18 patients with chronic venous leg ulcers were cultured. Six of these patients had F. magna. None of these F. magna strains had any hydrolytic enzyme activity or affected the endothelial cell proliferation. All inhibited fibroblast proliferation and keratinocyte wound repopulation. Björk18 was the first to describe a novel bacterial cell-wall protein that is able to bind with Ig light chains (L chains); therefore, this protein was named protein L. L chains are

407

shared between the different Ig classes. Protein L was found to have affinity with IgG, IgM, IgA, F(ab’)2, and Fab fragments, and with κ and λ L chains. The reaction with λ L chains is very weak compared with κ L chains. Nilson et al.34 described that protein L binds exclusively to the VL domain of Ig and not to the CL domain. This binding strongly depends on the three-dimensional structure of the VL domain, indicating that several sites of VL are involved. It requires the spatial proximity of the κI, κIII, and κIV light chain molecules. κ L chains represent 65% of human immunoglobulins, and of the entire κ chain population, κI, κIII, and κIV proteins represent 60%, 28%, and 2%, respectively.35 Protein L has five highly homologous domains that are involved in the binding of Ig. These domains interact with the framework regions of the VL domain.36 The strength of the binding of protein L with κ chains is less when compared with the binding of the complete Ig. The conformation resulting from the interaction between heavy and light chains in the Ig provides a more favorable binding site for protein L.37 The binding site of Ig is close to the antigen-binding site, but the interaction between protein L and Ig was not obstructed by occupation of the antigen-binding site.37,38 Åkerström et al.37 showed that protein L has no disulfide bond or a subunit structure, and that protein L has two non-Ig-binding fragments that were found to be unique. This was confirmed by Graille et al.39 A single protein L domain can react with the variable regions of κ L chains of two Fab molecules, in a sandwich fashion. The contact residues in the variable region are remote from the hypervariable loops. It was suggested that the two binding sites on protein L have a different affinity for Ig. In vivo experiments by Smith et al.40 showed that protein L prefers to target B cells. This is due to the interaction with Ig on the surface of these cells. This interaction strongly activates the B cells, which results in an upregulation of MHC-II and CD86. These surface molecules are important in initiating an antibody response. No specific binding of protein L with other splenocytes, like T cells and certain dendritic cell subsets, was observed. The activation of B cells also results in an increased expression of the target immunoglobulin. When mature B cells are exposed to protein L, a reduction of splenic marginal zone B cells and peritoneal B1 cells was observed.41 These two B-cell subsets are involved in the firstline immune response against foreign invaders. They have a high antigen-presenting capacity and secrete preferentially potentially protective natural IgM. B1 cells are located in the cavities of the body and are important in contributing to the production of natural antibodies and T-cell independent immune responses. Marginal zone B cells are located at the periphery of the splenic periarteriolar lymphoid sheath at the border of white and red pulp, and they are the first to encounter blood-borne antigens. The overall design of protein L is similar to those of protein A from staphylococci and protein G from streptococci, but the primary structure is different. When the amino acid compositions of these proteins are compared, protein L has a higher amount of glycine and a lower amount of lysine. No amino acid sequence homology was demonstrated between

408

these three proteins, apart from the carboxyl-terminal transmembrane region. Some similarity was seen between the W-region of protein G and the amino acid sequence of one of the tryptic peptides of protein L. It has been proposed that this W-region anchors protein G to the cell wall. Protein L is much smaller in size than protein G, which facilitates tissue penetration.38 One common feature between the three proteins is that they all possess multiple copies of Ig-binding domains. In each protein these domains are highly conserved.39 Since protein L is able to bind with all human Ig, it is also able to bind with the κ L chain of IgE. Since the binding of anti-IgE with the Fc portion of IgE stimulates the release of histamine from human basophils, it is possible that the binding of protein L with IgE also stimulates the release of histamine, which will trigger an inflammatory response. Histamine and tryptase are both involved in allergic reactions. This stimulation of basophils was described by Patella et al.19 The release of histamine is dependent on the concentration of protein L. The interaction with IgE present on the surface of basophils mediates the release of protein L. The stimulation by protein L on the basophils is greater than the stimulation by anti-IgE. Patella et al.19 also described the release of the preformed de novo-synthesized mediators leukotriene C4 (LTC4) from basophils, or PGD2 from human skin mast cells, both chemical mediators of human inflammatory cells. Genovese et al.42 described the release of histamine from human heart mast cells. They found a significant correlation between histamine release and tryptase release, and the release of LTC4. It is interesting to note that mast cells tend to accumulate at the site of a chronic infection.43 The release of de novo-synthesized mediators may contribute to the pathogenesis of the infecting strain. It is hypothesized that this may cause mycocardial damage in patients with bacterial infections.42 Protein L helps the bacteria to adhere to the wound surfaces. The covering of the bacterial cell wall by host proteins allows the bacteria to evade the immune response of the host.38 Protein L is expressed at the surface of ±10% of the F. magna strains.38,44 Some protein L molecules are released into the growth medium, but most molecules are associated with the cell wall. The protein L from the growth medium shows a considerable heterogeneity in size. This indicates proteolytic degradation of the released protein.37 Kastern et al.44 found that only F. magna strains that express protein L possess the protein L encoding gene. In nonexpressing protein L strains, this gene is not present rather than being downregulated. The features described for protein L may explain why protein L-expressing F. magna strains are more often associated with clinical infections than are nonexpressing strains. Kastern et al.44 determined the presence of protein L in 30 F. magna strains, all derived from clinical specimens. Four of these strains expressed protein L, and all of these four strains were isolated from women with bacterial vaginosis. The negative strains were from healthy women (n = 19), men (n = 4), and women with bacterial vaginosis (n = 3). These results indicate that there is a correlation between protein

Molecular Detection of Human Bacterial Pathogens

L-expressing F. magna strains and bacterial vaginosis. This was confirmed by de Château et al.29 They determined the presence of protein L and HSA-binding protein in 48 F. magna strains. Thirty of these strains were isolated from suppurative infections. One of them expressed protein L, and 16 strains (53%) were binding HSA. Eight of the 48 strains were isolated from patients with bacterial vaginosis. None of these strains showed HSA binding, and five were expressing protein L. The remaining ten strains of the original 48 were commensal strains, and none of them was expressing protein L or binding HSA. These results confirm the correlation between protein L-expressing F. magna strains and bacterial vaginosis. Furthermore, it also shows that F. magna strains isolated from localized suppurative infections preferentially express HSA‑binding protein. It is striking to notice that no strains were found that expressed protein L and HSA-binding protein in combination. This was also noticed by Ng et al.45 A total of 32 F. magna strains, from different origins and countries, were analyzed. Strains were found to be protein L-expressing, HSA-binding protein expressing, or expressing neither of them. No strains were found to express both proteins. Molecular typing of these strains showed that protein L and HSA-binding strains are associated with genotypic clusters. Recently, two other proteins that enhance the virulence of F. magna have been described, that is, a subtilisin-like proteinase (SufA) and a F. magna adhesion factor (FAF). FAF is expressed by more than 90% of the F. magna strains. This protein is cell-wall bound and can be released in the growth medium. The soluble form of FAF causes large protein aggregates, and the cell-wall bound FAF induces bacterial aggregation through protein-protein interactions between the FAF proteins of the different F. magna cells.22 FAF has the typical features of surface proteins of gram-positive bacteria; a C-terminal part with a cell wall spanning region, a membrane anchor, and an intracellular charged tail. F. magna strains which express FAF interact with BM40 and colonize the skin in the same way as SufA-expressing F. magna strains. Another feature of FAF is that it protects F. magna against LL-37, an antimicrobial peptide.22 BM40 is able to stimulate wound healing and it increases the albumin transport across the endothelium. The increase of albumin will stimulate PAB-expressing F. magna strains in their growth. SufA is the first described proteinase for F. magna.20 It is associated with the cell wall, but is also released in the growth medium. Most F. magna strains possess homologs of SufA. SufA is able to degrade the antibacterial peptides LL-37 and MIG/CXCL9. Thus F. magna enhances its own growth and can spread to commensal areas, where it is not present under normal conditions. Since SufA is a subtilisin, it first has to undergo autocatalytic maturation before it can be active. For full enzymatic activity, dimer formation of SufA is required. The antibacterial peptide MIG/CXCL9 binds with the CXCR3 receptor, which activates the G-protein.46 This receptor is expressed on eosinophils, NK cells, activated T cells, and endothelial cells. Remarkably, MIG/ CXCL9 degraded by SufA is still able to bind to the CXCR3

409

Finegoldia

receptor.21 The fragments of MIG/CXCL9 are still able to kill Streptococcus pyogenes, but F. magna remains unaffected. This may be explained by the fact that the dimer formation is affected due to the fragmentation of MIG/CXCL9. This may result in a reduced antibacterial activity against F. magna. During infections caused by S. pyogenes, an ecological niche can be created for F. magna. SufA is also able to release the FAF protein from the cell wall of F. magna. Karlsson et al.21 described that FAF is able to bind with MIG/CXCL9 with a high affinity. The release of FAF from the cell wall results in a decrease of the antibacterial activity of MIG/CXCL9, thereby promoting the growth of F. magna during inflammation. In human plasma, SufA degrades fibrinogen, a major clotting enzyme.47 It increases the thrombin-induced coagulation time in a dose-dependent manner. Fibrinogen is cleaved by thrombin to create fibrin, which forms a temporary matrix in which cells can proliferate during wound repair. In its soluble form SufA forms dimers and/or multimers, which are proteolytically more active when compared with the monomers.47 Fibrinogen consists of three pairs of nonidentical chains Aα, Bβ, and γ .48 First, SufA removes the C-terminal portion of fibrinogen Aα chains. Second, the NH2-terminal part of the Bβ chains is attacked. At higher concentrations of SufA, the Aα chains are further processed, thereby removing the central polymerization sites. SufA associated with the cell wall prevents the formation of fibrin networks by binding to keratinocytes.47 When the skin is damaged or infected, F. magna SufA-expressing bacterial cells will be in contact with plasma proteins. The fibrinogen present in the plasma will be broken down by SufA present on the cell wall. The formation of a fibrin network is delayed. The fibrinopeptides (FPA and FPB) that are released during the cleavage of fibrinogen are chemotactic agents for neutrophils, macrophages, and fibroblasts. They also exert antibacterial properties against gram-negative and gram-positive bacteria.49 It seems that besides the clotting, other fibrinogen-mediated processes are also disturbed by SufA. FAF- and SufA-expressing strains might impair wound healing, as has been described by Stephens et al.33

36.1.3  Genome Organization Recently, the genome of F. magna ATCC29328 was assessed by Goto et al.50 It consists of a circular chromosome (1.797.577 bp, average G + C content 32.3%) and a plasmid pPEP1 (189.163 bp, average G + C content 29.7%). Complete gene sets for the biosynthesis of glycine, serine, threonine, aspartate, and asparagine were present. There were no carbohydrate phosphotransferase system (PTS) genes present for glucose, maltose, mannose, glucitol, cellobiose, and lactose. PTS genes for mannitol, galactitol, and sucrose were incomplete. A lot of genes encoding aminoacid/oligopeptide transporters were found on the genome. This enables F. magna to take up amino acids from the environment for growth and survival. Genes for superoxide reductase, NADH oxidase, and putative NADH dehydrogenase were also present. They probably are important for the survival of F. magna

in intermediate aerobic conditions, such as mucosa and the skin. Virulence factors for antiphagocytosis were encoded by genes present on the chromosome and the plasmid, one on each. In total, four genes encoding albumin-binding protein homologs were present, three on the chromosome, and one on the plasmid. In total, 10 genes encoding collagen-binding proteins were found, five on the chromosome, and five on the plasmid. In total, 20 genes encoding N-acetylmuramoyl-lalanine amidase homologs (Cwp66) were encountered, most of them located on the chromosome. These proteins play a role in the adherence of bacteria to host cells. The presence of genes encoding sortase was assessed on the chromosome and plasmid. On the chromosome four genes encoding sortase, homologs were present, and on the plasmid, seven. The pre­ sence of seven sortase homologs on the plasmid is especially interesting. It is the highest number of sortase homologs present on a plasmid, for as far as genome sequences are determined for gram-positive bacteria. This feature might be unique for F. magna. Since plasmids are considered to be of foreign origin, the amount of sortase homologs present on a plasmid may play an important role in the pathogenesis of F. magna. Sortases are extracellular transpeptidases that catalyze the cell anchoring of cell-wall proteins. Sortases can be grouped into four or five different classes.51,52 Each subgroup has its own preference for substrates, depending on the amino acids present in the cell-wall sorting signal pentapeptide motif. Sortase A is the most important one and catalyzes the highest number of substrates. The precursor of a cell-wall-bound protein is synthesized in the cytoplasm with an N-terminal signal peptide and a C-terminal sorting signal (Figure 36.1). The cell-wall sorting signal consists out of a pentapeptide motif (for sortase A: LPXGT motif) a hydrophobic region, and a tail of charged residues.53 The N-terminal signal peptide directs the precursor to the membrane for translocation. It is assumed that, after cleavage by a signal peptidase, the hydrophobic region and the positively charged tail retain the precursor within the secretory pathway until the sortase has recognized the substrate.28 The membrane-anchored sortase A cleaves between the threonine and glycine residues of the LPXTG-like motif.54 An amide-bond is formed between the carboxyl group produced by the cleavage of the LPXGT motif and peptidoglycan of the cell wall. Since most virulence factors are displayed on the cell wall, sortases play an important role in the virulence of bacteria. The amount of cell-wall anchored protein will be enriched and more varied. This may result in an enhancement of interaction between host tissue and other bacteria in mixed infections. Therefore, sortases are a possible new target for the development of new therapeutic drugs against bacterial infections.55

36.1.4  Diagnosis Phenotypical Techniques. F. magna strains are difficult to identify, since the strains do not show any saccharolytic activity and only produce acetate as volatile fatty acids (VFAs). Identification is therefore based on negative reactions.

410

Molecular Detection of Human Bacterial Pathogens

NH2 Medium

4 NH2

NH2

NAM

NAG NAM

-Glu

-Ala

NH2 - (Gly)5 -Lys

-Glu

-Ala (Gly)5 -Lys

sortase

LP G XT

NH2 Signal peptide

LP

XT G

Cell wall sorting signal

-Ala -Glu

- (Gly) -Lys 5

-Ala (Gly)5

-Ala (Gly)5

L PXT

1

TNH

X

-Ala

NAM LP

3

2

NAG

NAG

G

Cell wall

+

+

Cytoplasm

+

FIGURE 36.1  A schematic overview of the mechanism of covalent binding of surface proteins. (Adapted from Cossart, P., and Jonquières, R., Proc. Natl. Acad. Sci. USA, 97, 5013, 2000 [Copyright 2000 National Academy of Sciences, USA].) 1. The precursor protein is directed by the N-terminal region of the signal peptide to the cell membrane, after which it is cleaved by a signal peptidase (arrow). 2.  The precursor retains within the secretory pathway, probably due to the positively charged tail and the hydrophobic region. 3. The pentapeptide motif LPXTG is cleaved by sortase between the threonine and glycine residues. An amide-bond is formed between the produced carboxyl and peptidoglycan in the cell wall. 4.  The surface protein is anchored in the cell wall.

Microscopic appearance may be rather variable and therefore is not suitable for identification. Ezaki et al.56 showed that GPAC can easily be identified by their amidase and oligopeptidase activities. Murdoch et al.7 characterized the described species in further detail, among them F. magna. In total, nine reference strains of F. magna and 78 clinical isolates were analyzed. All strains showed similar proteolytic activity. Strongly positive reactions were obtained for arginine, leucine, and pyroglutamyl aminopeptidase. Negative reactions were obtained for proline, phenylalanine, and glutamylglutamyl aminopeptidases. Variations were observed for other proteolytic enzymes. No acid production from mannose, raffinose, glucose, and trehalose was found. A minor part of the strains (22%) was able to produce a small amount of acid from fructose. Most of the strains were negative for alkaline phosphatase, but 7% was found to be positive, including the type strain. Nineteen percent of the strains were able to produce catalase. All strains were resistant to sodium polyanethol sulfonate (SPS). If a metronidazole disk is used to exclude anaerobic staphylococci or capnophilic streptococci, it should be noted that F. magna is able to produce colonies in the inhibition zone after 48 hours of incubation. Recently, Song et al.57 developed a flow chart that made the identification of GPAC easier. In this flow chart, Parvimonas micra

and F. magna are differentiated from the other GPAC by their inability to produce β-glucuronidase and to ferment glucose and their ability to produce pyroglutamyl arylamidase. P. micra and F. magna are differentiated from each other by the production of proline arylamidase; P. micra is a producer and F. magna is not.57 Two current methods to phenotypically identify anaerobes are the Vitek system and Matrix-Assisted Laser Desorption Ionisation Time of Flight Mass Spectometry (MALDITOF-MS). Recently, BioMérieux (Marcy, France) has developed a new colorimetric identification card (ANC card), which can be used in the Vitek 2 system for the identification of anaerobes, including F. magna. Evaluation of this card showed that F. magna can be reliably identified using this method.58,59 MALDITOF-MS is a promising new method for identifying bacteria; however, no evaluation studies have been published yet that describe the suitability of the method for the identification of GPAC, including F. magna. Genotypical Techniques. Nowadays, nucleic acid–based techniques are available to improve the identification of bacteria. Song et al.60 evaluated if 16S rRNA sequencing is suitable for the identification of GPAC. They established that the quality of the sequences in the public databases can be

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poor and may lead to misidentification. They sequenced the type strain of F. magna and 36 clinical isolates. When the sequences of the clinical isolates were compared with the sequenced type strain, the homology was >99%. However, when they were compared with the sequences available in the public databases, homology was 1000 organisms per sample.14 Since the late 1990s, NAATs have become the preferred methods for detection of C. trachomatis infection. NAATs provide enhanced sensitivity and noninvasive diagnostic/ screening options. For example, the possibility of testing urine samples or self-collected vaginal swabs, which facilitates otherwise cumbersome screening. Today, NAATs are recommended for all types of urogenital samples except in cases with medico-legal implications such as rape or sexual abuse, where additionally culture of C. trachomatis is mandatory.46 The fundamental feature of NAATs is their ability to multiply and, in principle, detect down to a single copy of the target DNA or RNA. However, the detection of a nucleic acid is not to be taken as a sign of a current infection. A positive NAAT could reflect a variety of states, including (i) a current clinically active infection, (ii) the presence of residual DNA from a previous infection, (iii) the presence of residual DNA as a result of stochastic or systematic contamination in the laboratory, or (iv) a genuine false-positive result.47 NAATs may therefore have a lower positive predictive value for detecting current infection than generally expected. Test-of-cure may be indicated when symptoms persist in patients treated appropriately with antibiotics. NAATs detect nucleic acids from dead C. trachomatis and may lead to

Molecular Detection of Human Bacterial Pathogens

false-positive results in a cured patient within 4 weeks after completion of treatment. Culture or a nonamplification assay should therefore be used as test-of-cure within that period, bearing in mind that these assays may miss a persistent infection because they often are less sensitive than NAATs. Persistent symptoms may also be caused by untreated concomitantly sexually transmitted pathogens (e.g., Neisseria gonorrhoeae and Mycoplasma genitalium). 45.1.3.1  Conventional Techniques Cell Culture. Historically, cell culture has been the reference standard for detection of C. trachomatis. A breakthrough in the detection of chlamydial infections by culture was made in 1965, when Gordon and Quan published their paper on the use of irradiated McCoy cells for isolation of the Trachoma Agent in cell culture.48 This culture technique was simple in comparison to the existing isolation of microorganisms from embryonated hen eggs. The use of the irradiated McCoy cell system and a sensitive, further simplified technique using cycloheximide-treated McCoy cells49 made it possible to screen for Chlamydia in the cervix and fallopian tubes of patients with acute salpingitis.50 A disadvantage of cell culture is, however, the potentially suboptimal sensitivity in areas where fast transportation of samples and adequate culture techniques/facilities cannot be met. The main advantage of cell culture is that the specificity is 100% per definition. This “eliminates” the risk of false-positive results, which is of major concern when low-prevalence populations are tested or when medico-legal aspects are involved. Today, nucleic acid based methods have, in general, replaced cell culture. Though, if optimal transport and culture techniques/facilities are used, cell culture may be as sensitive as NAATs.51 Direct Microscopy. Until the beginning of the 1980s, the routine diagnosis of chlamydial infection was based on the demonstration by direct microscopy of pathognomonic cell inclusions of cell cultures inoculated with clinical material. Chlamydial inclusions were initially demonstrated, either by simple staining with iodine (C. trachomatis only) or Giemsa stain. Direct microscopy of EBs in clinical specimens became possible when the monoclonal antibody technology facilitated the use of Immunofluorescence in the early 1980s. The advantage was that DFA did not depend on viable C. trachomatis particles and a rapid cold transport chain. A disadvantage was, however, the need of fluorescence microscopy equipment, and careful staff skilled in microscopy. The labor-intensive and eye-tiring methodology precluded a more comprehensive screening for C. trachomatis in clinical samples. This became possible by the use of monoclonal antibodies and the introduction of the enzyme immunoassay technique. Serology. Serology is of limited value in the diagnosis of current chlamydial infection. One problem is that systemic and local antibodies may persist for years after the infection has resolved. Another problem was the low sensitivity and specificity of earlier serology assays based on whole chlamydial antigens. Chlamydial antibodies have, however, been

515

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useful for demonstrating associations between C. trachomatis and clinical conditions such as ectopic pregnancy and tubal factor infertility.52 Antibodies directed against epitopes on MOMP encoded by the omp1 gene divide C. trachomatis strains into 15 serovars A–L53 (Table 45.1). The distribution of serovars varies, but in general, serovar E seems to be dominating in most countries.53 Among men who have sex with men, certain subtypes of serovars D, G, and J seem to predominate.54 The serovars L1–L3 are associated with Lymphogranuloma venereum and have previously been mainly confined to developing countries. In recent years, a spread in Europe and North America of serovar L2 among men who have sex with men has been observed, albeit with a somewhat changed symptomatology.55 The commercially available Enzyme Immunoassays (EIAs) all use the LPS as antigen. Cross-reacting organisms such as strains of Acinetobacter calcoaceticus, Escherichia coli, Gardnerella vaginalis, Neisseria gonorrhoeae, and group B streptococci make the EIA assays less suitable for samples taken from anatomical sites with an extensive normal flora (e.g., the pharynx and rectum).56,57 Diagnostic performance (specificity) can be improved by testing positive samples by another assay (confirmatory test).58 The analytical sensitivity of EIA has been improved by signal amplification.59 Rapid enzyme immunoassay-based diagnostic tests for chlamydial antigen were developed for point-of-care testing in the doctor’s surgery. A recent study on an internetavailable C. trachomatis test for home use showed an assay yielding more false-positive (18/193) than true-positive (7/38) results.60 45.1.3.2  Molecular Techniques Molecular detection assays can be separated into nonamplified DNA hybridization (probe) tests and amplification tests.14 In probe tests, an oligonucleotide sequence (probe) hybridizes to a target within bacterial DNA or RNA present in the clinical specimens, and a signal is detected by Chemiluminescence. The NAAT assays utilize oligonucleotide sequences (primers) to hybridize to and flank the target in the genomic DNA or RNA and then multiply exponentially the target sequence by various molecular techniques.

Probe Tests. The PACE2 assay from Gen-Probe is a probe test targeting ribosomal RNA (rRNA) of C. trachomatis. The Gen-Probe PACE2 assay uses the intrinsic “amplification” of the target rRNA molecule, which is present in hundreds of copies per Chlamydia cell. The Digene Hybrid Capture II probe test uses RNA probes that recognize and hybridize with specific DNA sequences of C. trachomatis. The Digene test has been cleared for use on an automated instrument. The probe tests are being phased out in favor of NAATs. NAATs. Genomic or plasmid DNA sequences or rRNA are the main targets for molecular detection of C. trachomatis. Plasmid-free C. trachomatis isolates have occasionally been reported,30,61 but they seem to be a negligible problem. However, the new C. trachomatis variant with a 377 bp deletion in the target region for the Abbott and Roche assays62 has led to modifications of the two assays using a dual target approach (Table 45.2). Ribosomal RNA is used as target in the Gen-Probe Aptima Combo 2 assay (23S rRNA), the GenProbe single analyte APTIMA C. trachomatis assay (16S rRNA). The advantage of using ribosomal RNA molecules is the presence of targets in high numbers in the chlamydial cell leading to an enhanced analytical sensitivity. Furthermore, since ribosomal RNA molecules are among the most conserved sequences, the likelihood of genetic changes leading to false-negative results should be minimal. In-house C. trachomatis PCRs based on genes encoding 16S rRNA have also been described.63,64 Real-time PCR allows for the quantification of bacterial load. Michel et al.65 evaluated the C. trachomatis load in matched specimens from different anatomic sites. For men bacterial load was highest for urine (1200 EBs per 100 µL) and significantly lower for urethral swabs (821 EBs per 100 µL). The mean organism loads for infected women were 2,231, 773, 162, and 47 EBs per 100 µL for endocervical swabs, self-collected vaginal swabs, urethral swabs, and first void urine samples, respectively. NAAT assays offer a reliable method for the use of noninvasive samples such as urine (males) and self-collected vulvo-vaginal specimens, which makes larger screening programs possible. Genotyping. Serotyping and genotyping based on single genes of C. trachomatis have only been of limited success in the studies of C. trachomatis epidemiology. Genotype

TABLE 45.2 Targets for Selected Commercial C. trachomatis NAAT Assays Company and Assay Abbott RealTime CT new formulation Becton Dickinson ProbeTec™ CT Gen-Probe APTIMA COMBO 2® Gen-Probe APTIMA® CT COBAS® TaqMan® CT Test, v2.0 Siemens VERSANT™ kPCR CT a

NAAT Principle Real-time PCR (Dual plasmid targets) Strand Displacement Amplification Transcription Mediated Amplification Transcription Mediated Amplification Real-time PCR (Dual targets) Real-time PCR

Within the deletion area of the Swedish plasmid mutant. ORF, Open reading frame.

Sequence within Target Specified A: ORF 1a; B: ORF 3 ORF 4 23S rRNA molecule 16S rRNA molecule A: ORF 1a; B: chromosomal omp1 gene ORF 2

516

based on PCR and sequencing of omp1 coincides with the MOMP serotype, and genotyping is now replacing serotyping. The subject was recently reviewed by Pedersen et al.53 Typing of C. trachomatis may be wanted for several purposes. A major aspect is the analysis of transmission patterns in sexual networks, and to look for possible association between types and specific disease entities or syndromes. An interesting but unresolved issue is to discriminate between persistence and reinfection in persons with repeatedly positive Chlamydia tests. Genotyping may also be crucial in investigations of sexual assaults. However, developing highly discriminating genotyping methods is difficult because of the highly conserved genome of C. trachomatis. With the aim of analyzing evolutional changes, a Multilocus sequence typing (MLST) system for the entire Chlamydiaecae family based on seven housekeeping genes was developed and could discriminate between reference strains of C. trachomatis, but provided almost no resolution for Cp. pneumoniae.66 Recently, two alternative detection systems have been reported to provide high resolution typing of C. trachomatis, and of other Chlamydiaceae species. High resolution MLST typing of C. trachomatis strains has been achieved by sequencing five target regions67; analysis of 47 clinical isolates of representative serotypes resulted in 32 genetic variants among 12 omp1 variants. An alternative multiplelocus variable-number tandem repeat analysis (MLVA) system for high resolution genotyping of C. trachomatis describes an analysis of variable numbers of tandem repeats (VNTR) in three loci combined with omp1 sequencing that reached a significantly higher diversity index than omp1 alone.68 The combined omp1 and VNTR genotyping system was readily performed on samples submitted in the GenProbe transport media for the Aptima Combo 2 assay for C. trachomatis.

45.2  METHODS Three main issues need to be addressed before the selection of a molecular Chlamydia assay: target population; specimen types and handling (collection and transport); and the practical implementation of the assay. Before implementation of an assay, the laboratory should consider handling of false-positive results, control of sample inhibition and contamination, quality control of the assay, and reporting (results and/or interpretations). An adequate number of epithelial cells infected with C. trachomatis in the specimen collected is crucial for any assay. No test, however, is better than the weakest link in the assay procedure. For NAATs this comprises sample preparation (extraction and purification of nucleic acids), amplification, and detection.

45.2.1  Sample Preparation Urine in particular but also other specimen types may contain inhibitors of the NAATs. Nucleic acid extraction methods,

Molecular Detection of Human Bacterial Pathogens

therefore, play an important role in the overall performance of an assay. The extraction can be generic (i.e., purifying all nucleic acids in the sample), or target specific (i.e., purifying nucleic acids containing the target of the assay only). Use of target capture thus increases the specificity of the particular assay but the generic methods provide purified material for other NAAT assays. Automated extraction methods generally apply magnetic particles to which the nucleic acids are bound by various principles. A magnet retains the particles in the sample tube during the washing procedures whereas potential inhibitors and other substances are removed. Urogenital Samples. All commercial NAAT assays are FDA approved for urogenital swabs. Vaginal swab specimens are the preferred specimen type for chlamydial screening of females. Vaginal swab specimens are as clinical sensitive and specific as cervical swab specimens.69 Furthermore, a vaginal swab, as with a urine sample may be self-collected at home or in the STD clinic or the GPs surgery. Cervical samples are still preferred by most doctors when full pelvic examinations are carried out, but vaginal swab specimens may constitute an appropriate sample type. Urine samples combined with a cervical swab specimen have been shown to significantly improve the yield when compared with urine samples only.70 Liquid-based Pap screening for cervical cancer may also provide sample material for testing by FDA approved C. trachomatis assays.71 For males, a urine specimen is preferred for two reasons: (i) NAATs are at least as sensitive on urine as for a urethral swab72 and (ii) urine sampling is noninvasive, which may motivate more men to be examined. However, urine specimens must contain an adequate number of cells with C. trachomatis and hence a “first-void” urine sample should be collected preferably at a time when the last urination was no less than 1 h earlier. Extragenital Samples. The NAAT assays are not FDA approved for use on specimens from extragenital sites such as the eye, pharynx, and rectum. An in-house validation should therefore be performed if specimens from extragenital sites are tested. In the final report, the laboratory should state that the specimen is not approved for the assay. Furthermore, the result should be interpreted with caution, if an independent NAAT has not been used for confirmation. It is surprising that commercial companies have not validated their assays for pharyngeal and rectal swabs considering the potential market for these tests.

45.2.2  Detection Procedures All diagnostic tests should be evaluated for their analytic and clinical performance before routine use. The analytic specificity of a Chlamydia NAAT is determined by testing a variety of bacteria, parasites, fungi, viruses, yeast, and so forth that may be isolated from the urogenital tract or other sites of sampling. The analytic sensitivity is determined by testing a serial dilution of all serovars of C. trachomatis. The clinical validity of a test is a measure of its ability to detect the associated disease or disorder. This may be the difficult part

Chlamydia

due to the lack of a clinical standard for Chlamydia infection, especially when no clinical signs or symptoms are present. 45.2.2.1  Commercial Assays Manufacturers of commercial NAAT assays detecting C. trachomatis have different NAAT principles, targets, and hardware solutions (Table 45.2). Real-time PCR is used by Abbott, Roche and Siemens, Strand Displacement Amplification (SDA) by Becton Dickinson, and Transcription Mediated Amplification (TMA) by Gen-Probe. Choice of assay platform depends on a series of practical and economical factors of which the performance can be scrutinized through External Quality Assessment (EQA) programs. These are supplied by, for example, the College of American Pathologists or by European organizations such as United Kingdom National External Quality Assessment Service (UK NEQAS), and Quality Control for Molecular Diagnostics (QCMD). In surveys or reports from these EQA programs, the rate of false-positive and false-negative results according to the product manufacturer can be compared. 45.2.2.2  Combo-Testing Dual detection of the both C. trachomatis and N. gonorrhoeae in the same urogenital sample has been FDA approved for the major commercially available NAATs. However, the advantage of the combo-testing may be outweighed by its inappropriate use in testing populations with a low prevalence of for example, gonorrhea. Even with an acceptable specificity of the N. gonorrhoeae assay, the number of false-positive tests may be unacceptably high. The BD Viper™ and the TIGRIS® DTS™ System from Gen-Probe both offer a high throughput automated platform for combined testing capable of analyzing more than 350 samples for both C. trachomatis and N. gonorrhoeae per 8-h shift. Compared to manual platforms these robots significantly reduce manual labor and repetitive movements for the laboratory staff. These two platforms were evaluated and compared with the more recent Abbott RealTime PCR system m2000,73 which has a lower throughput (186 specimens per 8-h shift). It was concluded that all three assays were suitable for the detection of the two microorganisms.73 45.2.2.3  S ensitivity and Specificity of Commercial NAATs A systematic review assessing the sensitivity and specificity of commercial NAATs for detection of C. trachomatis in urine and cervical (women) and urethral (men) swab samples, respectively, was published by Cook et al. in 2005.74 The sensitivities were between 83.3% and 96.7% with specificities in the range 93.8%–99.6%. Assays based on TMA and SDA found significantly more C. trachomatis than PCR assays in cervix specimens. In female urine, TMA found significantly more C. trachomatis than SDA. For male samples (urethra and urine), no significant difference in sensitivity was observed for PCR, SDA, and TMA. Results of TMA and PCR were nearly identical for urine and cervix specimens, whereas the sensitivity of SDA for C. trachomatis in cervix and urine specimens was significantly different.

517

Notably, with a decreasing prevalence of the Chlamydia disease, even specificity figures >99% may create increasing problems with false-positive tests and a decreasing positive predictive value (PPV) of the assay. With a disease prevalence of 1%, the false-positive rate (1-PPV) of an assay with 90% sensitivity and 99% specificity is then more than 50%. 45.2.2.4  In-House PCR Assays In-house or “home-brewed” PCR tests for Chlamydia can be useful as a supplement to the commercial C. trachomatis assay or as the only option when a commercial assay is not available (e.g., in detection of other Chlamydia species inclu­ ding Cp. Pneumoniae).75–78 Using a second target, in-house PCRs may serve as confirmatory testing of samples with equivocal test results in commercial assays. In-house assays may also be used for identification of plasmid-free strains61 or the new variant of C. trachomatis that was discovered in Sweden.79 There are also several important applications for in-house PCR assays where methods for subspeciation or genotyping are wanted53 or when the plasmids and genomes of the various Chlamydia species are compared.28 A specific real-time in-house PCR test for detection of C. trachomatis serovar L1–L3 causing Lymphogranuloma venereum has been described.80 Principle.  C. trachomatis can be identified and genotyped by PCR amplification and sequencing analysis of the omp1 gene53 and three VNTRs68 using the primers shown in the Table 45.3. Procedure 1. To amplify the omp1 gene, prepare PCR mixture (50 µL) comprising 5 µL Extensor Hi-fidelity PCR Mix 10× (Thermo Fisher), 0.8 µL dNTP (25 mmol of each), 1 µL (100 pmol) each of forward and reverse primers, 0.25 µL of Extensor polymerase, 36.95 µL H2O, and 5 µL of extracted DNA. The primers used for generating an approximately 1.1-kb fragment of the omp1 gene are PCTM3 (F) and NRI (R). 2. The thermo cycler program is as follows: 120 s at 94°C, followed by a touch-down protocol of 5 cycles of (denaturation at 94°C for 10 s, annealing at 65°C–1°C/cycle for 30 s, extension at 68°C for 60 s), 45 cycles of (10 s at 94°C, 30 s at 60°C, and 60 s at 68°C), and finally one cycle of 7 min at 68°C. 3. To amplify all three VNTR genes, prepare a multiplex PCR mixture (50 µL) consisting of 5 µL AmpliTaq Gold® buffer 10× (Applied Biosystems), 10 µL of MgCl 25mM, 0.8 µL dNTP (25 mmol of each), 1 µL (100 pmol) of each forward (CT1291 F, CT1335 F) and reverse (CT1291 F, CT1335 F) primers, and 0.25 µL each of the CT1299 F and CT1299 R primers, 24.2 µL H2O, 0.5 µL AmpliTaq Gold® polymerase, and 5 µL of extracted DNA 4. The thermo cycler program is 10 min at 94°C, 40 cycles of (45 s at 94°C, 45 s at 56°C, and 45 s at 72°C), and finally one cycle of 10 min at 72°C.

518

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TABLE 45.3 Primers for Identification and Sequence Analysis of C. trachomatis Primer PCTM3 NRI CT1291 F CT1291 R CT1299 F CT1299 R CT1335 F CT1335 R a





Sequence (5′→3′)

Positiona

TCCTTGCAAGCTCTGCCTGTGGGGAATCCT CCGCAAGATTTTCTAGATTTC GCCAAGAAAAACATGCTGGT AGGATATTTCCCTCAGTTATTCG TTGTGTAAAGAGGGTCTATCTCCA AAGTCCACGTTGTCATTGTACG TCATAAAAGTTAAATGAAGAGGGACT TAATCTTGGCTGGGGATTCA

58–84 1073–1053 195, 536–195, 555 195, 760–195, 738 291, 758–291, 781 291, 945–291, 924 737, 225–737, 250 737, 377–737, 358

Specificity Omp1 gene Hypothetical protein CT172.1 Noncoding region DNA topoisomerase

According to the omp1 nucleotide sequence of C. trachomatis serovar strain D/UW-3/Cx.11

5. Electrophorese the products on a 2% (w/v) agarose gel, and visualize with ethidium bromide stain. 6. Purify PCR products before sequencing is performed. Excise the band of the expected size from the 2% agarose gel, and purify DNA using the QIAquick Gel Extraction Kit (QIAgen). An insertion mutation may have occurred in the VNTR CT1291 region since in some strains the band is 224, and in others approximately 510 bp. 7. Perform sequencing reactions with the BigDye® v 3.1 Cycle Sequencing Kit (Applied Biosystems) according to the manufacturer’s instructions using 6 μL master mix (2 μL BigDye, 0.4 μL (10 pmol/μL) primer (F or R), 3.6 μL H2O) and 4 μL template. The thermo cycler program is 120 s at 94°C, 25 cycles of 10 s at 94°C, 5 s at 50°C, and 4 min at 60°C, and hold at 8°C. Sequence the omp1 once with the forward and once with the reverse primer, creating an overlap of approximately 250 bp in the mid region. Genotype the omp1 gene sequences using the program standard nucleotide-nucleotide Basic Local Alignment Search Tool (BLAST) (http://www.ncbi.nlm.nih. gov/BLAST/) “BLAST two sequences” approach. Use accession numbers for the omp1 reference types listed in paper by Pedersen et al.68 Investigate visually the sequences of the VNTRs (only the repeated part of the sequences) for differences.68

45.2.2.5  Quality Control Essentially, quality control is performed to secure that assays do not produce false-negative and false-positive results (retain highest possible sensitivity and specificity). To ensure good laboratory practice, the performance of NAATs should be scrutinized by a series of internal and external controls. Failure to follow the regulations and requirements for performance, monitoring, auditing, recording, analysis, and reporting of test results can be devastating to patients and laboratory. An internal certification should be performed even if an approved commercial test is implemented. This can be done by use of preselected panels of test samples with known

results or initially by running the new assay in parallel with the old routine system, bearing in mind that a new more sensitive assay may require confirmatory testing. Use of an in-house assay necessitates a more thorough validation. The analytical sensitivity should be calculated by determining the limit of detection through serial dilutions of known amounts of purified DNA and cultured bacteria spiked into the relevant specimen matrix. Spiked samples should be purified by the new sample extraction procedure before testing. The analytical specificity of the in-house assay should be checked with isolates covering a broad geographical and temporal spectrum, as well as different patient populations. Internal controls are useful in the daily runs. Correct test results for the positive and negative controls monitor the general performance of the assay. An internal amplification control (IAC) (i.e., a small amount of a known target) may be added to each reaction in order to demonstrate lack of inhibition in the individual clinical sample.81 The IAC should be added before the NA extraction procedure in order to also control for accidental loss of the specimen during purification. Lack of amplification of the IAC may also document partial reagent or instrument failure affecting only a subset of the specimens. Periodically, the laboratory can perform an internal quality assessment by selecting a panel of specimens with known results and relabel them before repeat testing as for ordinary clinical specimens. Confirmatory testing of positive results has been recommended, although the procedure may be unnecessary.82 There are three supplemental testing approaches: (i) repeating the original test on the original specimen, (ii) retesting the original specimen with a different NAAT, and (iii) performing a different test on a duplicate specimen. However, as pointed out by Hadgu and Sternberg,83 if a test result cannot be confirmed by itself, it is not important whether it is confirmed or not by another test. On the other hand, failure to repeat positive results could reflect the variability inherent with low copy numbers of targets. Use of external controls (blind samples) is an important quality control, and laboratories should participate in organized EQA schemes. It is essential that the quality control samples are tested according to the standard method used in

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the laboratory including all pre- and postexamination procedures. Exchanging specimens between laboratories using different NAAT methodologies may also be of help in validation of NAAT results. Last but not least, laboratories should provide guidelines for proper sampling and educate health care personnel to ensure correct use of the assays and interpretation of results.





45.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES Detection methods based on nucleic acid amplification led to Chlamydia being detected in novel tissues and cells, for example, human joints, atherosclerotic plaques, brains, and amoebae. With improved molecular identification and typing methods for Chlamydia, new insight into host-parasite relations and spread of these pathogens can be envisaged. The epidemiology of C. trachomatis infection in young people worldwide indicates that we still have more to learn about preventing the spread of this pathogen and the sequelae of its infection. We may also have to reevaluate the meaning of (e.g., a positive NAAT assay in asymptomatic patients and consider whether this result reflects true infection with C. trachomatis and whether the patient needs to be treated with antibiotics). Induction of protective immunity by vaccination against C. trachomatis may hopefully constitute a future option in the prevention of sexually transmitted infections with Chlamydia. Today, cell culture detection of Chlamydia is rarely used. This may, in combination with the lack of nonculture assays for drug susceptibility test prevent disclosure of inadequate antimicrobial treatment of asymptomatic infections. Drug resistant C. trachomatis strains have been described,84 but they do not at present seem to be a widespread clinical problem. However, the rapid spread of a mutant variant of C. trachomatis in Sweden62 speaks for surveillance also for drug resistant mutants. Future developments are likely to include multiplex STD panels combined with detection systems based upon a single detection chip or a liquid array technology. By creating multiplex PCR detection systems that not only detect all relevant microorganisms but also determine their drug resistance pattern and genotype, we may have concomitant information for treatment and prevention of sexually transmitted infections.

REFERENCES

1. Collier, L.H., Recent advances in the virology of trachoma, inclusion conjunctivitis and allied diseases, Br. Med. Bull., 15, 231, 1959. 2. Tang, F.F. et al., Studies on the etiology of trachoma with special reference to isolation of the virus in chick embryo, Chin. Med. J., 75, 429, 1957. 3. Page, L.A., Revision of the family Chlamydiaceae Rake (Rickettsiales): Unification of the PsittacosisLymphogranuloma venereum-Trachoma group of organisms in the genus Chlamydia Jones, Rake and Stearns, 1945, Int. J. Syst. Bacteriol., 16, 223, 1966.







4. Wagar, E.A., Safarians, S., and Pang, M., Analysis of genomic DNA from Chlamydia trachomatis for Dam and Dcm methylation, FEMS Microbiol. Lett., 77, 161, 1992. 5. Storz, J., and Page, L.A., Taxonomy of the Chlamydiae: Reasons for classifying organisms of the genus Chlamydia, family Chlamydiaceae, in a separate order, Chlamydiales ord. nov., Int. J. Syst. Bacteriol., 21, 1971. 6. Everett, K.D., Bush, R.M., and Andersen, A.A., Emended description of the order Chlamydiales, proposal of Parachlamydiaceae fam. nov. and Simkaniaceae fam. nov., each containing one monotypic genus, revised taxonomy of the family Chlamydiaceae, including a new genus and five new species, and standards for the identification of organisms, Int. J. Syst. Bacteriol., 49, 415, 1999. 7. Read, T.D. et al., Genome sequence of Chlamydophila caviae (Chlamydia psittaci GPIC): Examining the role of nichespecific genes in the evolution of the Chlamydiaceae, Nucleic Acids Res., 31, 2134, 2003. 8. Stephens, R.S. et al., Divergence without difference: Phylogenetics and taxonomy of Chlamydia resolved, FEMS Immunol. Med. Microbiol., 55, 115, 2009. 9. Fox, A. et al., Muramic acid is not detectable in Chlamydia psittaci or Chlamydia trachomatis by gas chromatographymass spectrometry, Infect. Immun., 58, 835, 1990. 10. Stephens, R.S. et al., Genome sequence of an obligate intracellular pathogen of humans: Chlamydia trachomatis, Science, 282, 754, 1998. 11. Skipp, P. et al., Shotgun proteomic analysis of Chlamydia trachomatis, Proteomics, 5, 1558, 2005. 12. Barbour, A.G. et al., Chlamydia trachomatis has penicillinbinding proteins but not detectable muramic acid, J. Bacteriol., 151, 420, 1982. 13. Montigiani, S. et al., Genomic approach for analysis of surface proteins in Chlamydia pneumoniae, Infect. Immun., 70, 368, 2002. 14. Black, C.M., Current methods of laboratory diagnosis of Chlamydia trachomatis infections, Clin. Microbiol. Rev., 10, 160, 1997. 15. Caldwell, H.D., Kromhout, J., and Schachter, J., Purification and partial characterization of the major outer membrane protein of Chlamydia trachomatis, Infect. Immun., 31, 1161, 1981. 16. Caldwell, H.D., and Schachter, J., Antigenic analysis of the major outer membrane protein of Chlamydia spp., Infect. Immun., 35, 1024, 1982. 17. Wang, S.P. et al., Immunotyping of Chlamydia trachomatis with monoclonal antibodies, J. Infect. Dis., 152, 791, 1985. 18. Kiselev, A.O. et al., Expression, processing, and localization of PmpD of Chlamydia trachomatis serovar L2 during the chlamydial developmental cycle, PLoS One, 2, e568, 2007. 19. Stothard, D.R., Toth, G.A., and Batteiger, B.E., Polymorphic membrane protein H has evolved in parallel with the three disease-causing groups of Chlamydia trachomatis, Infect. Immun., 71, 1200, 2003. 20. Kalman, S. et al., Comparative genomes of Chlamydia pneumoniae and C. trachomatis, Nat. Genet., 21, 385, 1999. 21. Read, T.D. et al., Genome sequences of Chlamydia trachomatis MoPn and Chlamydia pneumoniae AR39, Nucleic Acids Res., 28, 1397, 2000. 22. Carlson, J.H. et al., Comparative genomic analysis of Chlamydia trachomatis oculotropic and genitotropic strains, Infect. Immun., 73, 6407, 2005. 23. Kari, L. et al., Pathogenic diversity among Chlamydia trachomatis ocular strains in nonhuman primates is affected by subtle genomic variations, J. Infect. Dis., 197, 449, 2008.

520 24. Caldwell, H.D. et al., Polymorphisms in Chlamydia trachomatis tryptophan synthase genes differentiate between genital and ocular isolates, J. Clin. Invest., 111, 1757, 2003. 25. Palmer, L., and Falkow, S., A common plasmid of Chlamydia trachomatis, Plasmid, 16, 52, 1986. 26. Sriprakash, K.S., and Macavoy, E.S., Characterization and sequence of a plasmid from the trachoma biovar of Chlamydia trachomatis, Plasmid, 18, 205, 1987. 27. Thomas, N.S. et al., Plasmid diversity in Chlamydia, Microbiology, 143, 1847, 1997. 28. Seth-Smith, H.M. et al., Co-evolution of genomes and plasmids within Chlamydia trachomatis and the emergence in Sweden of a new variant strain, BMC Genomics, 10, 239, 2009. 29. Stothard, D.R. et al., Identification of a Chlamydia trachomatis serovar E urogenital isolate which lacks the cryptic plasmid, Infect. Immun., 66, 6010, 1998. 30. An, Q. et al., Infection with a plasmid-free variant Chlamydia related to Chlamydia trachomatis identified by using multiple assays for nucleic acid detection, J. Clin. Microbiol., 30, 2814, 1992. 31. Peterson, E.M. et al., The 7.5-kb plasmid present in Chlamydia trachomatis is not essential for the growth of this microorganism, Plasmid, 23, 144, 1990. 32. Carlson, J.H. et al., Comparative genomic analysis of Chlamydia trachomatis oculotropic and genitotropic strains, Infect. Immun., 73, 6407, 2005. 33. Ripa, T., and Nilsson, P., A variant of Chlamydia trachomatis with deletion in cryptic plasmid: Implications for use of PCR diagnostic tests, Euro Surveill., 11, E061109.2, 2006. 34. Møller, J.K., Pedersen, L.N., and Persson, K., Comparison of Gen-probe transcription-mediated amplification, Abbott PCR, and Roche PCR assays for detection of wild-type and mutant plasmid strains of Chlamydia trachomatis in Sweden, J. Clin. Microbiol., 46, 3892, 2008. 35. Lusher, M., Storey, C.C., and Richmond, S.J., Plasmid diversity within the genus Chlamydia, J. Gen. Microbiol., 135, 1145, 1989. 36. Tam, J.E. et al., Location of the origin of replication for the 7.5kb Chlamydia trachomatis plasmid, Plasmid, 27, 231, 1992. 37. Fenton, K.A., and Lowndes, C.M., Recent trends in the epidemiology of sexually transmitted infections in the European Union, Sex. Transm. Infect., 80, 255, 2004. 38. Østergaard, L. et al., Home sampling versus conventional swab sampling for screening of Chlamydia trachomatis in women: A cluster-randomized 1-year follow-up study, Clin. Infect. Dis., 31, 951, 2000. 39. Richardus, J.H., and Götz, H.M., Risk selection and targeted interventions in community-based control of chlamydia, Curr. Opin. Infect. Dis., 20, 60, 2007. 40. Scholes, D. et al., Prevention of pelvic inflammatory disease by screening for cervical chlamydial infection, N. Engl. J. Med., 334, 1362, 1996. 41. Kasi, P.M. et al., Blinding trachoma: A disease of poverty, PLoS Med., 1, e44, 2004. 42. Cates, W. Jr., and Wasserheit, J.N., Genital chlamydial infections: Epidemiology and reproductive sequelae, Am. J. Obstet. Gynecol., 164, 1771, 1991. 43. Klint, M. et al., Lymphogranuloma venereum prevalence in Sweden among men who have sex with men and characterization of Chlamydia trachomatis ompA genotypes, J. Clin. Microbiol., 44, 4066, 2006. 44. Rockey, D.D., Lenart, J., and Stephens, R.S., Genome sequencing and our understanding of Chlamydiae, Infect. Immun., 68, 5473, 2000.

Molecular Detection of Human Bacterial Pathogens 45. Stephens, R.S., The cellular paradigm of chlamydial pathogenesis, Trends Microbiol., 11, 44, 2003. 46. Centers for Disease Control and Prevention, Workowski, K.A., and Berman, S.M., Sexually transmitted diseases treatment guidelines, 2006, MMWR Recomm Rep., 55, 1, 2006. 47. Hadgu, A., Issues in Chlamydia trachomatis testing by nucleic acid amplification test, J. Infect. Dis., 193, 1335, 2006. 48. Gordon, F.B., and Quan, A.L., Isolation of the trachoma agent in cell culture, Proc. Soc. Exp. Biol. Med., 118, 354, 1965. 49. Ripa, K.T., and Mårdh, P.A., Cultivation of Chlamydia trachomatis in cycloheximide-treated mccoy cells, J. Clin. Microbiol., 6, 328, 1977. 50. Mårdh, P.A. et al., Chlamydia trachomatis infection in patients with acute salpingitis, N. Engl. J. Med., 296, 1377, 1977. 51. Møller, J.K., Østergaard, L.J., and Hansen, J.T., Clinical evaluation of four non-related techniques for detection of Chlamydia trachomatis in endocervical specimens obtained from a low prevalence population, Immunol. Infect. Dis., 4, 191, 1994. 52. Persson, K., The role of serology, antibiotic susceptibility testing and serovar determination in genital chlamydial infections, Best Pract. Res. Clin. Obstet. Gynaecol., 16, 801, 2002. 53. Pedersen, L.N., Herrmann, B., and Møller, J.K., Typing Chlamydia trachomatis: From egg yolk to nanotechnology, FEMS Immunol. Med. Microbiol., 55, 120, 2009. 54. Mossman, D. et al., Genotyping of urogenital Chlamydia trachomatis in Regional New South Wales, Australia, Sex. Transm. Dis., 35, 614, 2008. 55. van de Laar, M.J., The emergence of LGV in Western Europe: What do we know, what can we do? Euro Surveill., 11, 146, 2006. 56. Hammerschlag, M.R., Rettig, P.J., and Shields, M.E., False positive results with the use of chlamydial antigen detection tests in the evaluation of suspected sexual abuse in children, Pediatr. Infect. Dis. J., 7, 11, 1988. 57. Taylor-Robinson, D., Thomas, B.J., and Osborn, M.F., Evaluation of enzyme immunoassay (Chlamydiazyme) for detecting Chlamydia trachomatis in genital tract specimens, J. Clin. Pathol., 40, 194, 1987. 58. Østergaard, L., and Møller, J.K., Use of PCR and direct immunofluorescence microscopy for confirmation of results obtained by Syva MicroTrak Chlamydia enzyme immunoassay, J. Clin. Microbiol., 33, 2620, 1995. 59. Okadome, A. et al., Reactivity of a dual amplified chlamydia immunoassay with different serovars of Chlamydia trachomatis, Int. J. STD AIDS, 10, 460, 1999. 60. Michel, C.E. et al., Pitfalls of internet-accessible diagnostic tests: Inadequate performance of a CE-marked Chlamydia test for home use, Sex. Transm. Infect., 85, 187, 2009. 61. Magbanua, J.P. et al., Chlamydia trachomatis variant not detected by plasmid based nucleic acid amplification tests: Molecular characterisation and failure of single dose azithromycin, Sex. Transm. Infect., 83, 339, 2007. 62. Ripa, T., and Nilsson, P.A., A Chlamydia trachomatis strain with a 377-bp deletion in the cryptic plasmid causing falsenegative nucleic acid amplification tests, Sex. Transm. Dis., 34, 255, 2007. 63. Mahony, J.B. et al., Evaluation of the NucliSens Basic Kit for detection of Chlamydia trachomatis and Neisseria gonorrhoeae in genital tract specimens using nucleic acid sequencebased amplification of 16S rRNA, J. Clin. Microbiol., 39, 1429, 2001.

Chlamydia 64. Madico, G. et al., Touchdown enzyme time release-PCR for detection and identification of Chlamydia trachomatis, C. pneumoniae, and C. psittaci using the 16S and 16S-23S spacer rRNA genes, J. Clin. Microbiol., 38, 1085, 2000. 65. Michel, C.E. et al., Chlamydia trachomatis load at matched anatomic sites: Implications for screening strategies, J. Clin. Microbiol., 45, 1395, 2007. 66. Pannekoek, Y. et al., Multi locus sequence typing of Chlamydiales: Clonal groupings within the obligate intracellular bacteria Chlamydia trachomatis, BMC Microbiol., 8, 42, 2008. 67. Klint, M. et al., High-resolution genotyping of Chlamydia trachomatis strains by multilocus sequence analysis, J. Clin. Microbiol., 45, 1410, 2007. 68. Pedersen, L.N., Pødenphant, L., and Møller, J.K., Highly discriminative genotyping of Chlamydia trachomatis using omp1 and a set of variable number tandem repeats, Clin. Microbiol. Infect., 14, 644, 2008. 69. Schachter, J. et al., Vaginal swabs are the specimens of choice when screening for Chlamydia trachomatis and Neisseria gonorrhoeae: Results from a multicenter evaluation of the APTIMA assays for both infections, Sex. Transm. Dis., 32, 725, 2005. 70. Airell, A. et al., Chlamydia trachomatis PCR (Cobas Amplicor) in women: Endocervical specimen transported in a specimen of urine versus endocervical and urethral specimens in 2-SP medium versus urine specimen only, Int. J. STD AIDS, 11, 651, 2000. 71. Chernesky, M. et al., Abilities of APTIMA, AMPLICOR, and ProbeTec assays to detect Chlamydia trachomatis and Neisseria gonorrhoeae in PreservCyt ThinPrep Liquid-based Pap samples, J. Clin. Microbiol., 45, 2355, 2007. 72. Sugunendran, H. et al., Comparison of urine, first and second endourethral swabs for PCR based detection of genital Chlamydia trachomatis infection in male patients, Sex. Transm. Infect., 77, 423, 2001. 73. Levett, P.N. et al., Evaluation of three automated nucleic acid amplification systems for detection of Chlamydia trachomatis and Neisseria gonorrhoeae in first-void urine specimens, J. Clin. Microbiol., 46, 2109, 2008.

521 74. Cook, R.L. et al., Systematic review: noninvasive testing for Chlamydia trachomatis and Neisseria gonorrhoeae, Ann. Intern. Med., 142, 914, 2005. 75. Mitchell, S.L. et al., Evaluation of two real-time PCR chemistries for the detection of Chlamydophila pneumoniae in clinical specimens, Mol. Cell. Probes, 23, 309, 2009. 76. Robertson, T. et al., Characterization of Chlamydiaceae species using PCR and high resolution melt curve analysis of the 16S rRNA gene, J. Appl. Microbiol., 107, 2017, 2009. 77. Tang, Y.W. et al., Qualitative and quantitative detection of Chlamydophila pneumoniae DNA in cerebrospinal fluid from multiple sclerosis patients and controls, PLoS One, 4, e5200, 2009. 78. McKechnie, M.L. et al., Simultaneous identification of 14 genital microorganisms in urine by use of a multiplex PCRbased reverse line blot assay, J. Clin. Microbiol., 47, 1871, 2009. 79. Catsburg, A. et al., TaqMan assay for Swedish Chlamydia trachomatis variant, Emerg. Infect. Dis., 13, 1432, 2007. 80. Morre, S.A. et al., Lymphogranuloma venereum diagnostics: From culture to real-time quadriplex polymerase chain reaction, Sex. Transm. Infect., 84, 252, 2008. 81. Hoorfar, J. et al., Practical considerations in design of internal amplification controls for diagnostic PCR assays, J. Clin. Microbiol., 42, 1863, 2004. 82. Moncada, J., Donegan, E., and Schachter, J., Evaluation of CDC-recommended approaches for confirmatory testing of positive Neisseria gonorrhoeae nucleic acid amplification test results, J. Clin. Microbiol., 46, 1614, 2008. 83. Hadgu, A., and Sternberg, M., Reproducibility and specificity concerns associated with nucleic acid amplification tests for detecting Chlamydia trachomatis, Eur. J. Clin. Microbiol. Infect. Dis., 28, 9, 2009. 84. Somani, J. et al., Multiple drug-resistant Chlamydia trachomatis associated with clinical treatment failure, J. Infect. Dis., 181, 1421, 2000.

46 Chlamydophila Chengming Wang, Bernhard Kaltenboeck, and Konrad Sachse CONTENTS 46.1 Introduction...................................................................................................................................................................... 523 46.1.1 Classification, Morphology, and Biology............................................................................................................. 523 46.1.2 Clinical Features and Pathogenesis...................................................................................................................... 524 46.1.3 Diagnosis.............................................................................................................................................................. 526 46.1.3.1 Conventional Techniques....................................................................................................................... 526 46.1.3.2 Molecular Techniques............................................................................................................................ 527 46.2 Methods............................................................................................................................................................................ 529 46.2.1 Sample Preparation............................................................................................................................................... 529 46.2.2 Detection Procedures............................................................................................................................................ 529 46.2.2.1 Real-Time PCR (TaqMan®) Detection of Chlamydiaceae and Chlamydophila spp. ........................... 529 46.2.2.2 Real-Time FRET PCR (LightCycler®) Detection of Chlamydiaceae spp. ........................................... 530 46.2.2.3 Detection of Chlamydophila spp. Using the ArrayTube® Test...............................................................531 46.3 Conclusion and Future Perspectives..................................................................................................................................531 References.................................................................................................................................................................................. 532

46.1  INTRODUCTION 46.1.1  Classification, Morphology, and Biology The traditional classification of chlamydiae was mainly based on host susceptibility and the biological properties of the pathogens.1,2 But in the past three decades, molecular phylogenetic investigations have revolutionized our view of the taxonomic relationships of microorganisms.3–5 Studies of phylogenetic relationships between the major outer membrane protein (ompA) genes of chlamydiae laid the groundwork for the classification based on genetic relatedness.6 Further phylogenetic analyses of other chlamydial genes, most notably the ribosomal RNA genes, demonstrated essentially identical evolutionary relationships among chlamydiae. A subsequent classification scheme, based on the sequence differences of the 16S and 23S ribosomal RNA genes, separated the family Chlamydiaceae into two genera, Chlamydia and Chlamydophila7–10 (Figure 46.1). The reclassification of Chlamydiaceae into two genera did not meet with wide approval from scientific colleagues, public health workers, and funding agencies.11,12 Therefore, reclassification of Chlamydiaceae into a single genus Chlamydia has been proposed, containing nine species named by the current species epithets.12 The Chlamydiales are obligate intracellular bacterial pathogens of higher cells that infect eukaryotic cells, from single-celled organisms such as amoebae to virtually any cell type of multicellular organisms. The chlamydial elementary

body (EB) is the infectious stage of the chlamydial developmental cycle, and functions as a tough “spore-like” body whose purpose is to permit chlamydial survival in the nonsupportive environment outside the host cell. The EB is near the limit of light microscopic visibility being approximately 0.3 µm in diameter and round or, occasionally pear-shaped, containing electron-dense structures made up of tightly coiled genomic DNA covered by histone-like proteins. The ultrastructure of EBs has been extensively studied,13 and spikes in the membrane are thought to be the tubular apparatus of a chlamydial type III secretion system. The chlamydial reticulate body (RB) represents the noninfectious intracellular developmental stage with a diameter of approximately 1 µm. The RB is metabolically active and the cytoplasm is rich in ribosomes, which are required for protein synthesis. RBs replicate exclusively in an intracellular vacuole termed an “inclusion.” As the RBs begin to differentiate into EBs at the end of the chlamydial developmental cycle, sites of recondensation of nucleic acid appear in the inclusion’s cytoplasm. In the maturing inclusion, chlamy­ dial particles appear to be packed tightly within the inclusion membrane. Development of chlamydiae is highly dependent on nutrient supply and the metabolic status of host cells. Nutrient deficiencies such as low glucose, iron, or amino acid levels lead to delayed development and aberrant chlamydial organisms within the inclusion. These aberrant inclusions are thought to represent persistent chlamydial forms that are important for maintaining the infection in a host population, particularly in the absence of overt clinical signs. 523

524

Molecular Detection of Human Bacterial Pathogens

78

50 nt

100 97

100

Chlamydophila psittaci Chlamydophila caviae Chlamydophila felis

Chlamydophila

Chlamydophila pneumoniae

43

100

Chlamydophila abortus

Chlamydiaceae

Chlamydophila pecorum Chlamydia trachomatis

99

Chlamydia suis

100

Chlamydia

100 Chlamydia muridarum Waddliaceae

WSU 86-1044 56

Parachlamydia acanthamoebae Simkania negevesis

Parachlamydiaceae Simkaniaceae

FIGURE 46.1  Phylogeny of the order Chlamydiales based on full-length 16S rRNA genes.7,9 Branch lengths are measured in nucleotide substitutions, and numbers show branching percentages in bootstrap replicates. Only bacteria of the family Chlamydiaceae have fully established pathogenic potential for vertebrates. Strains of the remaining families Waddliacae, Parachlamydiaceae, and Simkaniaceae have been variably and inconsistently associated with diseases.

While sharing the unique biphasic developmental lifecycle, Chlamydophila spp. demonstrate assorted epidemiological characteristics. C. pneumoniae, first described in 1986,14 is mainly a human pathogen but also infects koalas, horses, and frogs.15–18 This pathogen is distributed worldwide and is arguably among the most common human pathogens. Acute, chronic, and asymptomatic infections with C. pneumoniae occur with high frequency in virtually all humans during their lifetime.19,20 The 5- to 20-year age group shows the most rapid rise in prevalence.19 C. pneumoniae contributes to 10% or more of the respiratory diseases in young adults, and to more serious respiratory disease in senior individuals. Kleemola et  al.21 reported epidemic outbreaks of C. pneumoniae infection among military recruits in Finland between 1957 and 1985. Each outbreak lasted approximately 6 months, and epidemics occurred during all seasons of the year. The infection rate varied from 60 to 80 per 1000 men. Transmission was slow, with the case-to-case interval averaging 30 days and ranging as high as 3 months.21 It is possible that all avian species are the natural hosts for C. psittaci as subclinical carriers.22,23 Stress factors such as crowding and shipping enhance shedding of the organisms in birds, resulting in high prevalence of the infection. Fatal human C. psittaci infection was first described as “psittacosis” in Switzerland in the 1870s, and it attracted more public attention in 1929 and 1930 during a widespread outbreak.24 Since most avian species are subclinical carriers of C. psittaci, persons with continual contact with birds are at great risk for psittacosis. This includes poultry farmers and processors, zoo workers, owners of pet shops and of racing pigeons, and laboratory workers.25,26 More recently, an outbreak of psittacosis in a poorly managed poultry flock involving two different genotypes of C. psittaci spread to about 100 small flocks in the surrounding region and led to

infection of 24 individuals, with one lethal case.27 Laroucau et al.28 reported five severe cases of psittacosis in individuals associated with duck farms in France in 2006, where human samples and duck isolates exhibited identical genetic characteristics in their PCR–RFLP restriction patterns. C. abortus causes infectious abortion and mastitis in sheep, goat, and cattle, as well as pneumonia in pigeons, turkeys, and sparrows.24 Abortion in sheep (ovine enzootic abortion, [OEA]) caused by C. abortus was first described by Grieg in Scotland in 1936,29 and chlamydial abortion has subsequently been recognized as one of the most important causes of abortion in sheep. Infection with C. abortus has also been reported to be associated with abortion in women.30,31 In most cases of C. abortus-induced human abortion, direct contact to infected sheep or goats occurred. The outcome of human infection in the first trimester of pregnancy is likely to be spontaneous abortion, whereas later infection causes stillbirths or preterm labor.32 Therefore, pregnant women must avoid all contact with lambing ewes and newborn lambs, and should not handle contaminated clothing from those working with these animals. Immunocompromised individuals also must avoid contact with potential sources of infection at lambing time.30

46.1.2  Clinical Features and Pathogenesis C. pneumoniae is primarily a human pathogen of the respiratory tract and causes acute or chronic bronchitis and pneumonia.33 Infections are mainly asymptomatic and unrecognized, or mildly symptomatic illnesses. The most frequently recognized disease manifestations associated with C. pneumoniae infections are pneumonia and bronchitis. In adults, 10% of cases of pneumonia and approximately 5% of bronchitis and sinusitis cases have been attributed to the organism.19,33 Severe

Chlamydophila

systemic infections with C. pneumoniae, while uncommon, do occur. The onset of disease is gradual, and illness is more frequent in young adults and older debilitated individuals.24 The incubation period of the symptoms may extend over several weeks. In these cases, pharyngitis and laryngitis precede lower respiratory disease. If lower respiratory tract symptoms occur, radiography usually demonstrates pneumonia. Numerous studies have demonstrated strong links between C. pneumoniae infection and obstructive pulmonary disease, metabolic syndrome, insulin resistance, Alzheimer’s disease, and cardiovascular disease.19,20,34–41 However, antibiotic prevention in large clinical trials, including a 1-year course of weekly azithromycin administration in a 4012-patient trial, failed to reduce secondary coronary events.42,43 In an obese mouse model, Wang et al.41 examined the longitudinal progression of insulin resistance and diabetes under influence of C. pneumoniae infection, genetic background, and dietary fat concentration. They concluded that murine C. pneumoniae infection enhances insulin resistance and diabetes in a genetically nutritionally restricted manner only in obese C57BL/6, but not A/J mice, via circulating inflammatory mediators such as TNF-α.41 C. psittaci primarily infects birds, but is readily transmissible to humans. C. psittaci infections in birds are often systemic with intermittent shedding of the organism, and may take a clinically inapparent, chronic, or acute course. Most organs, such as conjunctiva and the respiratory and gastrointestinal tracts, become infected. Stress will commonly trigger the onset of severe symptoms, and result in the rapid deterioration and death of the animals. Until recently, nine serovars of C. psittaci were distinguished, all of which were defined by epitopes on variable domains of the major outer membrane protein OmpA.44,45 Seven serovars were thought to predominantly occur in a particular order or class of Aves and two in nonavian hosts, that is, type A in psittacine birds; B in pigeons; C in ducks and geese; D in turkeys; E in pigeons, ducks, and others; E/B in ducks; F in parakeets; WC in cattle; and M56 in rodents. However, comprehensive analysis of all available ompA gene sequences revealed the limitations of this classification46 and the relative character of the assumed host specificity.46,47 Consequently, adjustments to the typing scheme were suggested, which include the introduction of subgroups to the more heterogeneous genotypes A, E/B, and D, as well as six new genotypes representing so-far untypable strains.46 Most of the avian genotypes have also been identified sporadically in cases of zoonotic transmission. Human psittacosis is relatively rarely diagnosed in the United States and Europe, but cases with mild symptoms may often be overlooked when species-specific testing for C. psittaci is not conducted. Most cases are associated with infected birds, and parrots or parakeets are the most common source of infections. The infection is transmitted to humans by the respiratory route during contact with sick birds or from the contaminated environment, particularly by inhalation of aerosolized dried bird feces. Human case-to-case transmission is very rare. Nevertheless, in a remarkable episode of

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psittacosis in the Bayou region of Louisiana, 18 individuals became infected by secondary or tertiary transmission from the wife of a trapper, and eight of them died from interstitial pneumonia.48 The incubation period for psittacosis ranges from 1 to 4 weeks. The initial symptoms may include malaise and pain in the limbs, and, more often, high fever, headache, and cough. The pulse rate is typically slow in relation to the temperature, and there is little or no evidence of chest consolidation. The spleen can be enlarged. Meningitis, meningoencephalitis, myocarditis, and endocarditis occur in severe cases, and the patients may become drowsy if the central system is affected.24 Patients may recover without treatment and the infection usually resolves within 2 to 3 weeks. Death, due to respiratory or cardiovascular failure, has been rare since the advent of antibiotics. Before the antibiotic era, the death rate was as high as 50% in psittacosis patients older than 50 years.24 Suppressing the avian sources of infection, such as import bans for psittacine birds during the 1929 psittacosis pandemic,49 effectively controls psittacosis. Similarly, tetracycline treatment of imported birds during a 30-day quarantine period, as practiced in the United States, effectively reduces C. psittaci shedding and controls psittacosis.24 C. abortus strains are widespread in ruminants, efficiently colonize the placenta, and are primarily associated with cases of abortion and weak neonates. It has been reported that human C. abortus infections can result from contact with infected sheep,50–52 goats,31,53,54 or cattle.55–57 The majority of these reports referred to abortions in pregnant sheep farmers, who were exposed to C. abortus during lambing. For example, in a Swiss farm, abortion affected 50% of the goats during the lambing season, and C. abortus was identified as the causing pathogen. During this time, a pregnant woman had contact with aborting goats, developed a severe generalized infection, and aborted herself. Her placenta contained C. abortus as shown by both immunohistochemistry and polymerase chain reaction (PCR).54 Thus, pregnant women should stay away from sheep, especially during lambing. Inhalation of infected material from sheep can also result in respiratory disease in nonpregnant humans, but the products of the sheep dairy industry are usually not a hazard to human health.24 C. pecorum is associated with polyarthritis, enteritis, pneumonia, and urogenital infections in cattle, sheep, goats, koala, and swine.58–60 C. felis was isolated from cat pneumonia in 1944 and is associated with pneumonia and conjunctivitis in cats that may occasionally be transmitted to owners, resulting in mild conjunctivitis.24,61–63 C. caviae was originally isolated from conjunctival scrapings of guinea pigs,64 and may also cause fertility disorders in these animals. A recent report has associated this species with rhinitis and conjunctivitis in horses.65 The disease outcome of chlamydial infection is determined by the interaction between chlamydial replication and the host response, and a series of pathogen- and host-associated factors.41,66 Different efficiencies in infectivity and replication of the pathogen determine consistent, but not absolute, associations between chlamydial strain, host, and disease

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manifestation. These bacteria produce only minimally toxic or nontoxic products and regulate their growth based on the availability of host cytoplasmic nutrients. The initial events that elicit the host’s innate immune response may determine the type of repair response and ensure the outcome of a chlamydial infection.66 Chronic granulomatous lesions of mononuclear cell aggregates and fibrosis are the most common disease manifestations for chlamydial infections. Repeated infection and host genetic susceptibility are the main factors that exacerbate chlamydial infections.41 Cell-mediated immunity plays an important role in immune protection as well as disease development of chlamydial infection, with CD4+ helper lymphocytes assuming the key role in a protective host response.67–69 Th1-type effector cytokines, such as IFN-γ, restrict chlamydial replication, contribute to a protective delayed-type hypersensitivity response,70,71 and exert the main proinflammatory effects via stimulation of the production of free radical molecules such as reactive oxygen (ROS) and reactive nitrogen oxide species (RNOS).72 These central inflammatory mechanisms play a pivotal role in pathogen elimination, but also in cellular signaling and regulation of the immune response to chlamydial infection.73,74 Ultimately, they are also the cause of disease by inducing “collateral” tissue damage in an excessive response to the typically minor threat of chlamydial infection. The macro- and micronutrient content of diets influences free radical production and outcomes of infectious diseases in rodent models.75–80 Wang et al.41 quantitatively dissected the immune and disease response to repeated C. pneumoniae lung infection by multivariate modeling at dichotomous levels of four effects: mouse strain (A/J or C57BL/6), dietary protein content (14% protein, 0.3% l-cysteine/0.9% l-arginine or 24% protein, 0.5% l-cysteine/2.0% l-arginine); dietary antioxidant content (90 IU α-tocopherol/kg vs. 450  IU α-tocopherol/kg and 0.1% g l-ascorbate), and time course (3 or 10 days postinfection). Contrast analyses indicated that chlamydial disease is characterized by a profound early T  helper cell suppression. Thus, the mechanisms that regulate the host T-cell population are the critical determinants of chlamydial pathogenesis.41 Still, we know very little about the mechanisms of the disease production by Chlamydophila infections. One of the major restrictions is that we do not yet have the genetic methodology to pursue the function of individual chlamydial genes.81 It is imperative to further explore the mechanisms of chlamydial diseases at the cellular and molecular levels of both pathogen and host.

46.1.3  Diagnosis 46.1.3.1  Conventional Techniques Since chlamydiae are obligate intracellular bacteria, tissue culture techniques are required for isolation and propagation. A number of different cell lines can be used, such as McCoy, Buffalo Green Monkey Kidney (BGMK), HeLa 229, and Hep-2.82–84 Embryonated hens’ eggs are also occasionally used, particularly when high yields are required. Cell culture

Molecular Detection of Human Bacterial Pathogens

is usually considered 100% specific, since the presence of chlamydial inclusions is an unambiguous demonstration of infection, but sensitivity is considerably lower (60%–80%) since not all strains are easy to culture. Results are available within 48–72 h for well-growing isolates, but may be delayed for several days in the case of more difficult samples. Infection of chlamydiae in cell culture can be enhanced by centrifugation and/or by chemical treatment of cultured cells, before or during infection, using for example cycloheximide. As isolation is necessary to demonstrate the viability of a field strain and also facilitates detailed characterization by molecular and biochemical methods, cell culture is still widely regarded as the gold standard in chlamydial diagnosis.85 However, diagnosis by cell culture requires very experienced laboratory personnel and strictly standardized protocols and, due to time and cost constraints, will remain a domain of specialized laboratories. The characteristic intracytoplasmic inclusions of chlamydiae can be visualized in smears and tissue preparations from pathological specimens using histochemical and immunohistochemical staining under light microscopy. Depending on the manifestation of the infection, typical samples include smears prepared from biopsies, bronchoalveolar lavage, and placental membranes, as well as swabs taken from nose, ear, pharynx, vagina, or the moist coats of aborted fetuses. Prepared smears can be stained for detection of chlamydiae using one of several staining procedures, for example, modified Machiavello, modified Giménez, Giemsa, or modified Ziehl-Neelsen stain.86,87 The latter is considered the most satisfactory method, where small coccoid EBs stain red/pink against a counterstained blue or green cellular background. However, while the procedure works well with heavily infected tissue, samples with low levels of infection can be easily overlooked. Furthermore, chlamydiae can be demonstrated in histological preparations using a variety of staining procedures. A simple method involves the histochemical staining of thin tissue sections (≤4 µm) with Giemsa after fixation in fluids such as Bouin and Carnoy.86 Dark-ground methylene blue staining, which has been shown to be a more reliable method for detecting C. abortus in fetal membranes than Giemsa, can also be used.88 However, both these techniques are nonspecific and can cross-react with other bacterial species. Therefore, care must be taken with interpretation of results. Immunohistochemical staining procedures that utilize monoclonal antibodies (mAbs) against chlamydial surface antigens, such as LPS or MOMP, are more sensitive in comparison to histochemical staining. A direct immunoperoxidase method for detecting C. abortus in formalin-fixed tissues has proved to be a very rapid and sensitive test.89 An alternative approach involves the use of a fluorescein-­conjugated antimouse antiserum in combination with the mAb. Enhanced labeling can be achieved using the more complex streptavidin-biotin method.90 All in all, immunoperoxidase staining of formalin-fixed, paraffin-embedded or cryostat tissue sections is commonly used for diagnostic purposes, as well as for epidemiological and pathogenesis studies.91–93

Chlamydophila

Most of the commercial enzyme immunoassays (EIAs) were designed for Chlamydia trachomatis. However, since they are based on the family-specific LPS antigen, in principle, they should also be suitable for detection of Chlamydophila spp., albeit with some qualification on performance. These assays include direct fluorescent antibody tests (e.g., IMAGEN, Celltech; Chlamydia-Direct IF, BioMerieux; Vet-IF, Cell Labs), plate-based ELISA (Chlamydiazyme, Abbott; IDEIA, Dako; IDEIA PCE, Dako; Pathfinder, Kallestad; ChlamydiaEIA, Pharmacia), and solid-phase ELISA (Clearview Chlamydia MF, Unipath; Surecell, Kodak). While a well-validated EIA can be a useful and easyto-handle diagnostic tool, many comparative studies have revealed limitations in terms of sensitivity and specificity.94–96 The performance of individual assays can depend on sample type and antigen load. Typical findings varied between 60% and 70% for both sensitivity and specificity as compared to culture and PCR. Generally speaking, labeled mAbs against chlamydial LPS for immunofluorescent microscopy and ELISAs kits using chlamydial LPS antigen work well, whereas capture ELISAs tend to show considerable proportions of false positives. The detection of specific antibodies can be conducted using the microimmunofluorescence (MIF) test.97 The fact that it utilizes purified elementary bodies as antigen facilitates identification of species-specific antibodies, although practical experience is required to differentiate the signal from family-specific reactions. The test was found to be useful in serological surveys, but a recent study raised serious doubts on its suitability for diagnosing human C. psittaci infection.98 The complement fixation test (CFT) was the first described LPS-based assay and, despite its rather complex setup and limited sensitivity, is still used in some laboratories. More recently developed antibody tests include the recombinant LPS-based rELISA (www.medac.de), which detects Chlamydiaceae-specific antibodies at higher sensitivity than CFT, and the recomLine Chlamydia IgG and IgA (www.mikrogen.de), a strip immunoassay allowing detection of antibodies to C. pneumoniae, C. psittaci, and Chlamydia trachomatis. The general significance of serology in diagnosis of acute infections is limited because chlamydial antibodies appear only after 7–10 days and early antibiotic therapy can suppress the regular humoral response. 46.1.3.2  Molecular Techniques Conventional DNA Amplification Methods. The broad use of the PCR and other DNA amplification methods, which began in the 1990s, revolutionized the diagnosis of infectious diseases by allowing rapid identification from clinical specimens and DNA-based inter- and intraspecies differentiation. The majority of published conventional PCR methods are based on targets in the ribosomal RNA operon7,8,99 or the ompA gene.100,101 An optimized nested PCR assay based on the elaborate primer system of Kaltenboeck et al.100 was modified and described in detail by Sachse and Hotzel.102 The first round of amplification generates a Chlamydiaceae-specific product,

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which serves as template in the second round, where speciesspecific primers are used. This assay was found to be very robust for routine use and proved the most sensitive among several protocols. Another well-validated assay for detection of Chlamydia trachomatis, Chlamydia pneumoniae, and Chlamydia psittaci (old nomenclature), which can also be run in triplex mode, was published by Reference 103. The high sensitivity of this procedure was attained through a touchdown enzyme time release methodology featuring the use of hotstart DNA polymerase, a touchdown protocol for annealing temperatures to improve primer binding specificity, and an enzyme time release protocol to allow 60 cycles to be run for improved sensitivity. Although a large number of PCR tests for chlamydiae have been published, it is not always clear from the paper whether the test has been properly validated. The use of poorly validated or unvalidated test protocols may lead to the generation of invalid data due to poor performance parameters of that test. This issue was addressed in a review by Apfalter et al.104 The authors emphasized that preanalytical procedures, sample preparation and DNA extraction, assay design and setup, as well as interpretation and confirmation of results should be subject to validation in order to ensure high accuracy of data and high specificity and sensitivity. If considering the use of a published amplification assay, the prospective user should always check the data for adherence to the general rules on test validation. Real-Time PCR. In addition to information on the presence or absence of a given pathogen, real-time PCR provides the possibility to quantitate the amount of the agent present in the sample. This can be achieved by cumulative measurement of the fluorescent signal generated by a specifically annealed dual-labeled fluorogenic probe upon exonuclease digestion.105,106 As a major advantage, real-time PCR does not require post-PCR sample handling, which precludes potential PCR product carryover contamination and results in more rapid and high-throughput assays. The procedure has a large dynamic range of target molecule detection compr­ ising at least five orders of magnitude. Quantitative real-time PCR is based on measurement in the log-linear phase. Fluorescence signals are generated by specific binding of dye molecules, such as SYBR Green I, to double-stranded DNA107 or by labeled oligonucleotide hybridization probes.105 As fluorescence readings of PCR in the log-linear phase can be correlated to the initial number of target gene copies, quantitation of the number of microbial cells present in the sample can be accomplished. Practically, DNA concentration (as copy number) is calculated from the number of amplification cycles necessary to generate a fluorescence signal of a given threshold intensity. A summary of published real-time PCR assays for chlamydiae is given in Table 46.1. Most of the family-specific real-time PCR tests are targeting the 23S rRNA gene. The methodologies developed by DeGraves et al.109 and Ehricht et  al.108 have been validated and used in routine testing. Details are described in Section 46.2.2. Detection limits were

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Molecular Detection of Human Bacterial Pathogens

TABLE 46.1 Real-Time PCR Methods for Chlamydiae Specificity

Target Gene

Chlamydiaceae Chlamydiaceae Chlamydiaceae, Chlamydia psittaci,a C. pneumoniae, C. pecorum, Chlamydia trachomatis C. psittaci C. psittaci C. psittaci C. abortus C. felis C. felis C. pecorum C. caviae Chlamydia suis

23S rRNA 23S rRNA 23S rRNA

Everett et al.7,8 Ehricht et al.108 DeGraves et al.109

ompA ompA incA ompA ompA ompA ompA ompA 23S rRNA

Geens et al.110,b Pantchev et al.111,c Menard et al.112 Pantchev et al.111,c Helps et al.113 Pantchev et al.114 Pantchev et al.114 Pantchev et al.114 Pantchev et al.114

a b c

Reference

Based on four-species classification of the family Chlamydiaceae. Includes primer sets for genotype-specific amplification. Includes internal amplification control.

reported to be in the order of 1 to 10 target copies. However, Ehricht et al.108 pointed out that the actual detection limit may be dependent on the integrity of target DNA. DeGraves et al.115 developed a highly sophisticated realtime PCR platform for sensitive detection of chlamydiae, which is distinguished by its optimized nucleic acid–extraction protocol ensuring high template yield, its step-down thermal cycling profile ensuring high product yield, and its design for the high-throughput regime of a diagnostic laboratory. This comprehensive and robust system is based on fluorescence resonance energy transfer (FRET) technology run on the LightCycler and allows detection of the pathogens both at genus and species level, according to the traditional four-species nomenclature. Geens et al.110 also used LightCycler technology to develop a panel of real-time PCR tests for C. psittaci. While the species-specific assay is based on SYBR Green detection and therefore is slightly less sensitive than fluorescent probe-based assays, the authors managed to design individual tests for the C. psittaci genotypes A, B, C, D, E, F, and E/B, respectively. Interestingly, some of the assays had to be designed as competitive reactions. As the closely related genotypes A, B, and E could not be distinguished by individual TaqMan probes, nonfluorescent competitor oligonucleotides had to be included alongside the probes to attain the necessary specificity. Using discriminatory target segments in the ompA gene and TaqMan technology, Pantchev et al.111,114 developed individual assays for C. psittaci, C. abortus, C. felis, C. caviae, C. pecorum, and Chlamydia suis (for details, see Section 46.2.2). In view of the close genetic relatedness between C. psittaci and C. abortus it was necessary to design a minor groovebinding (MGB™) probe. Furthermore, these assays include

an internal amplification control,116 a sensitive indicator of amplification efficiency and of the presence or absence of DNA polymerase inhibitors. Based on incA gene sequence analysis of five C. psittaci strains, Menard et al.112 published a TaqMan real-time PCR protocol for detection of this species from clinical samples. They also used a MGB probe to exclude cross-reactions with C. abortus. DNA Microarray Technology. The new possibilities provided by this methodological approach may be particularly beneficial for laboratory diagnosis of infectious diseases. This highly parallel approach allows DNA samples to be simultaneously examined by a large number of probes, which may be derived from a polymorphic gene segment and/or from different genomic regions. A specific microarray hybridization test can be regarded as an equivalent to resequencing the target site, so that DNA microarray-based tests can attain far higher discriminating power than PCR. Sachse et al.117 developed a microarray assay for the detection and differentiation of Chlamydia spp. and Chlamydophila spp. The test uses the commercially available ArrayTubeTM (AT) system (Clondiag Chip Technologies, www.clondiag. com), a less expensive system for processing low- and highdensity DNA arrays that involves spotted DNA chips of 3 × 3 mm size assembled onto the bottom of 1.5-mL plastic microreaction tubes. In contrast to other microarray equipment, hybridization and signal processing can be conducted in an easy and rapid fashion on standard laboratory equipment without additional devices, such as hybridization chambers. Hybridization signals are amplified by an enzyme-catalyzed precipitation reaction. A  CCD camera integrated in a light transmission reader is used to monitor DNA duplex formation by kinetic measurement of the precipitation reaction at each spot. Hybridization probes for all Chlamydiaceae spp. were designed on the basis of a multiple sequence alignment, from which a highly discriminatory segment in domain I of the 23S rRNA gene was identified. Several rounds of optimization and refinement led to the present version of the chip,118 which is carrying 28 species-specific probes (for all nine chlamydial species); three genus-specific probes for Chlamydia and Chlamydophila, respectively; five probes identifying the closest relatives Simkania negevensis and Waddlia chondrophila, as well as four hybridization controls (consensus probes) and a staining control (biotinylated oligonucleotide). The AT assay has been used for the direct detection of chlamydiae from clinical tissue as shown in a validation study.118 The sensitivity was shown to be equivalent to that of real-time PCR,108 thus rendering the test suitable for use in the diagnostic lab. Genotyping by Microarray. The great potential of the AT microarray technology for rapid intraspecies characterization has been demonstrated by the recent development of a genotyping assay for C. psittaci.46,83,117 This assay was shown to discriminate all established genotypes and to identify sofar untyped strains. Its high specificity, which allows detection of single-nucleotide polymorphisms, is due to the parallel

Chlamydophila

approach consisting in the use of 35 hybridization probes derived from VD2 and VD4 of the ompA gene. This new test represents a promising diagnostic tool for tracing epidemiological chains, exploring the dissemination of genotypes and identifying nontypical representatives of C. psittaci.

46.2  METHODS 46.2.1  Sample Preparation A large number of manufacturers offer DNA extraction kits for clinical samples from different matrices. To verify whether a particular kit is suitable for a given panel of specimens, it should be tested on a series of spiked samples containing defined numbers of Chlamydia cells. The first step with commercial kits is usually the lysis of bacterial and tissue cells using a special buffer (“lysis buffer”), the effectiveness of which is decisive for the kit’s performance. An optional RNase digestion is intended to remove cellular RNA. Subsequently, the lysate is centrifuged through a minicolumn, where the released DNA is selectively bound to a solid phase (modified silica, hydroxyl apatite, or special filter membrane). After washing, the DNA can be eluted with elution buffer or water. These DNA preparations are usually of sufficient purity and virtually free of DNA polymerase inhibitors. Large diagnostic laboratories will probably prefer to use robotic systems for DNA extraction. In the following paragraphs, we describe alternative DNA preparation protocols that proved satisfactory in our hands. They can be used if, for some reason, kits are not an option. Swabs (e.g., Nasal, Vaginal, Conjunctival), Mucus, Bronchoalveolar Lavage, and Sputum. The swab is placed into a 2-mL Safe-Lock tube containing 500 µL of lysis buffer (100 mM Tris-base, pH 8.5, 0.05% [v/v] Tween 20). The closed tube is vortexed thoroughly for 1 min and centrifuged at 12,000 × g for 30 s. To squeeze out the remaining liquid, the swab is placed into a 1-mL pipette tip whose lower half was cut off and transferred into a fresh tube. After centrifugation at 12,000 × g for 1 min, the liquid is combined with that in the first tube from the previous centrifugation. (In the case of mucus, bronchoalveolar lavage, or sputum samples, the previous steps are omitted.) The fluid or mucus is centrifuged at 12,000 × g for 15 min, the supernatant is discarded and the pellet resuspended in 50 µL of lysis buffer containing 20 µL of proteinase K (10 mg/mL in water). After incubation at 60°C for 2 h, proteinase K is inactivated by heating at 97°C for 15 min. Finally, the mix is centrifuged at 12,000 × g for 5 min to remove debris, and 5 µL of the supernatant is used as template for PCR. Tissue from Lung, Tonsils, Lymph Nodes, Spleen, Liver, and Other Organs. One hundred mg of homogenized tissue are boiled in 200 µL water in a plastic tube for 10 min. Subsequently, the tube is allowed to cool to room temperature. Optionally, proteinase digestion can be conducted to increase the final yield of DNA: 200 µL SDS (10 mg/mL) and 20 µL of proteinase K (10 mg/mL) are added and the mix is incubated at 55°C for 1 h. Following the addition of 200 µL

529

phenol (saturated solution in TE buffer), the vessel is vortexed vigorously for 1 min and centrifuged at 12,000 × g for 5 min. The (upper) aqueous phase is transferred into a fresh tube, 200 µL of chloroform-isoamyl alcohol (24:1, v/v) are added, and the mix is vortexed at highest intensity for 1 min. Again, the aqueous phase is pipetted into a fresh tube. To precipitate the DNA, 120 µL of isopropanol are added, the reagents are thoroughly mixed and incubated at room temperature for 10 min. Following centrifugation at 12,000 × g for 10 min, the supernatant is discarded, the DNA pellet is allowed to air dry for 30 min and redissolved in 20 µL TE buffer. One µL is used as template for an amplification reaction. Feces. One hundred mg of sample is mixed with 200 µL water by vortexing for 1 min. The following steps are identical to the protocol for tissue.

46.2.2  Detection Procedures 46.2.2.1  R  eal-Time PCR (TaqMan®) Detection of Chlamydiaceae and Chlamydophila spp. Principle.  The assays were designed as duplex amplification reactions, all of which include an internal amplification control (IAC). The latter was developed by Hoffmann et al.116 as a universal heterologous amplification control system for real-time PCR. It consists of a plasmid carrying a 712-bp fragment of the EGFP gene, which serves as the IAC template (Intype IC-DNA, Labordiagnostik, Leipzig), as well as primers EGFP1-F/EGFP10-R and probe EGFP-HEX. The oligonucleotide sequences are given in Table 46.2. In addition, a primer-probe system targeting β-actin can be used as an efficiency control of DNA extraction (to be run in duplex mode with the chlamydia PCR or in triplex including the IAC). Procedure.  The assays are performed in 96-well optical microtiter plates on an Mx3000P Real-Time PCR System (Stratagene). The final 25-µL reaction mixture contains 12.5  µL of 2× TaqMan Gene Expression Mastermix with ROX (Applied Biosystems), each primer at a final concentration of 0.3 µM, each probe of 0.2 µM, 2500 copies of the IAC template and 1 µL of sample DNA. The following cycling parameters are used: initial cycle of 95°C for 10 min; 45 cycles of 95°C for 15 s; and 60°C for 1 min. For quantitative determinations, each plate run should include decimal serial dilutions of purified genomic DNA from the respective chlamydial species containing 2 × 10 −1 to 2 × 105 IFU per PCR reaction, which are used as copy standards. The cycle threshold value (Ct) is calculated automatically by the MxPro4.01 software (Stratagene). Note: In a routine diagnostic setting, the Chlamydiaceaespecific real-time PCR assay has been successfully used as a screening test that is followed by the ArrayTube (AT) test (described in Section 46.2.2.3) to identify the chlamydial species involved,47,83 that is, samples are first examined by realtime PCR, and the positives are subjected to the AT assay.

530

Molecular Detection of Human Bacterial Pathogens

TABLE 46.2 Primers and Probes for Real-Time PCR Detection of Chlamydiae Specificity Chlamydiaceae

C. psittaci

C. abortus

C. pecorum

C. felis

C. caviae

Chlamydia suis

IAC

Chlamydiaceae

Amplicon Size (bp) Target Gene

Primers, Probe and Sequences (5′→3′) Ch23S-F: CTGAAACCAGTAGCTTATAAG CGGT Ch23S-R: ACCTCGCCGTTTAACTTAACTCC Ch23S-p: FAM-CTCATCATGCAAAAGGCACGCCG-TAMRA CppsOMP1-F: CACTATGTGGGAAGGTGCTTCA CppsOMP1-R: CTGCGCGGATGCTAATGG CppsOMP1-S: FAM-CGCTACTTGGTGTGAC-TAMRA (MGB® probe) CpaOMP1-F: GCAACTGACACTAAGTCGGCTACA CpaOMP1-R: ACAAGCATGTTCAATCGATAAGAGA CpaOMP1-S: FAM-TAAATACCACGAATGGCAAGTTGGTTTAGCG-TAMRA CppecOMP1-F: CCATGTGATCCTTGCGCTACT CppecOMP1-R: TGTCGAAAACATAATCTCCGTAAAAT CppecOMP1-S: FAM-TGCGACGCGATTAGCTTACGCGTAG-TAMRA CpfOMP1-F: TCGGATTGATTGGTCTTGCA CpfOMP1-R: GCTCTACAATGCCTTGAGAAATTTC CpfOMP1-S: FAM-ACTGATTTCGCCAATCAGCGTCCAA-TAMRA CpcavOMP1-F: GAATAACATAGCCTACGGCAAACATA CpcavOMP1-R: CGATCCCAAATGTTTAATGCTAAGA CpcavOMP1-S: FAM-CAAGATGCAGAATGGTCCACAAACGC-TAMRA Csuis23S-F: CCTGCCGAACTGAAACATCTTA Csuis23S-R: CCCTACAACCCCTCGCTTCT Csuis23S-S: FAM-CGAGCGAAAGGGGAAGAGCCTAAACC-TAMRA EGFP1-F: GAC CAC TAC CAG CAG AAC AC EGFP10-R: CTT GTA CAG CTC GTC CAT GC EGFP-HEX: HEX-AGC ACC CAG TCC GCC CTG AGC A-BHQ1 CPEC23SUP: GGG GTT GTA GGG TCG ATA ACG TGA GAT C CTR23SUP: GGG GTT GTA GGR TTG RGG AWA AAG GAT C CHL23SDN: GAG AGT GGT CTC CCC AGA TTC ARA CTA CHL23SLCR640: LCR640-CCT GAG TAG RRC TAG ACA CGT GAA AC-Phos CHL23SFLU: GRA YGA HAC AGG GTG ATA GTC CCG TA-6FAM

46.2.2.2  R  eal-Time FRET PCR (LightCycler®) Detection of Chlamydiaceae spp. Principle.  An integrated nucleic acid isolation and FRET PCR platform was developed to specifically detect, differentiate, and quantify all Chlamydiaceae spp. with high sensitivity.109,115 Sampling into guanidinium-based buffer for maximum preservation of nucleic acids and optimized nucleic acid extraction has vastly improved detection of low numbers of chlamydial genomes.109,115 Step-down thermal cycling and an excess of hot-start Taq polymerase vastly improved the robustness and sensitivity of the real-time PCR. The requirement for simultaneous side-by-side annealing of two FRET probes used for real-time detection of the amplification product ensures essentially 100% specificity. The amplification of Chlamydiaceae 23S rRNA allowed for the differentiation of chlamydial species and was more robust than amplification of the ompA gene at low target numbers. Procedure.  Total nucleic acid extraction is performed with the High-Pure PCR Template Preparation Kit® (Roche Molecular Biochemicals) by glass fiber matrix binding and low-volume elution in 40 µL total volume.109 The PCR buffer is 4.5 mM MgCl2, 50 mM KCl, 20 mM Tris-HCl, pH 8.4,

Reference

111

23S rRNA

Ehricht et al.108

76

ompA

Pantchev et al.111

82

ompA

Pantchev et al.111

76

ompA

Pantchev et al.114

78

ompA

Pantchev et al.114

84

ompA

Pantchev et al.114

118

23S rRNA

Pantchev et al.114

117

EGFP

Hoffmann et al.116

168

23S rRNA

DeGraves et al.109

supplemented with 0.05% each Tween 20 and Nonidet P-40, and 0.03% acetylated bovine serum albumin (Roche Molecular Biochemicals). Nucleotides are used at 0.2 mM (dATP, dCTP, dGTP) and 0.6 mM (dUTP). Primer CHL23SDN is used at 1 µM, primers CPEC23SUP and CTR23SUP at 0.5 µM each, the LightCycyler Red 640 probe CHL23SLCR640 at 0.2 µM, and the fluorescein probe CHL23SFLU at 0.1 µM. For each 20 µL total reaction volume, 1.5 U hot-start Platinum Taq DNA polymerase (Invitrogen) and 0.2 U heat-labile uracil-DNA glycosylase (Roche Molecular Biochemicals) are used.109 Thermal cycling consists of a 2 min at 95°C followed by 18 high-stringency step-down thermal cycles, 40 low-stringency fluorescence acquisition cycles, and melting curve determination between 50°C and 80°C. The thermal protocol for the 23S rRNA qPCR is 6 × 12 s @ 64°C, 8 s @ 72°C, 0 s @ 95°C; 9 × 12 s @ 62°C, 8 s @ 72°C, 0 s @ 95°C; 3 × 12 s @ 60°C, 8 s @ 72°C, 0 s @ 95°C; 40 × 8 s @ 54°C and fluorescence acquisition, 8 s @ 72°C, 0 s @ 95°C.109,115 Notes: In a routine diagnostic setting, the real-time FRET PCR (LightCycler®) platform109,115 has been successfully applied to quantify and differentiate the Chlamydiaceae spp. in clinical samples with high specificity and sensitivity.

Chlamydophila

46.2.2.3  D  etection of Chlamydophila spp. Using the ArrayTube® Test Principle.  Prior to the AT test, sample DNA is prepared by standard extraction protocols and amplified by a consensus PCR using a biotinylated primer (U23F-19: 5′-ATT GAM AGG CGA WGA AGG A-3′ and 23R-22: 5′-biotin-GCY TAC TAA GAT GTT TCA GTT C-3′). After hybridization and staining, unique species-specific hybridization patterns for all nine species of the family Chlamydiaceae are obtained and immediately processed by the Iconoclust software (Clondiag). Procedure. In biotinylation PCR, the reaction mix is composed of 1 µL DNA template, 0.2 µL (1 U) Taq DNA Polymerase (Fermentas), 2 µL 10× Taq Buffer with KCl, 1 µL MgCl2 (25 mM), 2 µL dNTP mix (1 mM each), 1 µL of each primer (0.5 µM, U23F-19 and 23R-22), and 12.8 µL deionized water. The sample DNA is amplified with 40 cycles of 94°C for 30 s, 55°C for 30 s, and 72°C for 30 s. The hybridization protocol starts with conditioning of the AT vessel by washing twice with 500 µL of Hybridization buffer 1 (Clondiag) at 30°C for 5 min. All incubations are conducted upon slight shaking (550 rpm) on a heatable horizontal tube shaker (Thermomixer comfort, Eppendorf). For denaturation, 1 µL of the biotinylated PCR product is diluted with 99 µL hybridization buffer in a separate tube, heated at 95°C for 5 min and put on ice. After transfer into the AT, hybridization is allowed to proceed at 58°C for 60  min. Subsequently, the supernatant is discarded, and the tube is washed with 500 µL 2 × SSC/0.01% Triton X-100 (40°C, 5 min), 500 µL 2 × SSC (30°C, 5 min) and 500 µL 0.2 × SSC (20°C, 5 min). Vacant binding sites of the microarray are blocked by incubation with a 2% solution of Blocking Reagent (Roche) in hybridization buffer at 30°C for 15 min. Subsequently, the AT is incubated with 100 µL of a 1:10,000 dilution of streptavidin-conjugated horseradish peroxidase (AT Staining Kit, Clondiag) followed by three-step washing as above. Finally, 100 µL of the peroxidase substrate (AT Staining Kit) is added. Immediately afterwards, measurement of the hybridization signal is started (at 25°C, using the ATR01 array tube reader), and the final image is recorded after 10 min of continuous precipitation. Processing of hybridization signals is done using the Iconoclust software (Clondiag). Note: In a routine diagnostic setting, the AT test has been successfully combined with the Chlamydiaceae-specific real-time PCR protocol described in Section 46.2.2.1,83 that is, samples are first examined by real-time PCR, and the positives are AT tested to identify the species involved.

46.3  C  ONCLUSION AND FUTURE PERSPECTIVES Chlamydophila spp. are intracellular pathogens of humans and animals that have evolved in the face of unique and complex long-term relationships with their hosts.2,24,119–121 Many pioneering chlamydia researchers have contributed to

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extraordinary advances in the last 20 years, such as robust detection by PCR, ligase chain reaction, as well as serology, improved animal models, and whole-genome sequencing of multiple chlamydial strains and species.81,109,122–125 This chapter mainly dealt with three human disease–related Chlamydophila species: C. pneumoniae, C. psittaci, and C. abortus. While C. pneumoniae is the dominant endemic human Chlamydophila species, C. psittaci and C. abortus also infect humans, breaking out from their respective endemically infected host populations. When diagnosed, infections by C. psittaci and C. abortus in humans are typically severe and sometimes lead to fatal diseases. However, tests for these infections are not routinely performed, and, therefore, the true prevalence and clinical picture are poorly understood. The availability of complete genome sequences of all chlamydial species has opened the possibility of sensitive monospecific ELISA tests for each species. Such tests will be required for a complete assessment of the prevalence of these infections in humans. In animals, recent diagnostic advances have dramatically increased detection frequencies of chlamydial infections.126–130 Thus, the paradigm has shifted from rare but severe Chlamydophila spp. infections to widespread, low-level endemic infections. While evidence concerning the effect on animal health of these ubiquitous clinically inapparent infections is just emerging, it remains incomplete. Most likely, these infections are associated with several production diseases that limit profits in agriculture, such as fertility disorders, increased somatic cell counts in milk (subclinical mastitis), and reduced weight gains and fitness of young animals.119,131,132 Similarly, in humans, the paradigm of C. pneumoniae infection has shifted from acute respiratory disease to that of a virtually ubiquitous, clinically inapparent infection that nevertheless is strongly epidemiologically linked to chronic inflammatory diseases, such as coronary atherosclerosis and metabolic syndrome.122 The current prevailing notion for the involvement of C. pneumoniae in atherosclerosis stipulates that C. pneumoniae-infected macrophages disseminate to early atherosclerotic lesions and exacerbate pathological processes.122 To explore these mechanisms, Wang et al.41 quantitatively examined, in an obese mouse model, the influence of C. pneumoniae infection on progression of insulin resistance, a central feature of metabolic syndrome that also underlies atherosclerosis. They demonstrated conclusively that low-copy C. pneumoniae organisms dispersed to secondary tissues are irrelevant to the progression of insulin resistance. Rather, acute infection of the lung, the primary target of C. pneumoniae, causes an increase in circulating cytokines that drive the long-term exacerbation of insulin resistance and accelerate onset of type 2 diabetes, metabolic syndrome, and presumably, atherosclerosis.41 The concept of circulating inflammatory mediators released from the main infection site is consistent with the mechanisms enhancing insulin resistance proposed for other prevalent pathogens that typically reside only at predilection infection sites, such as H. pylori,133 P. gingivalis,134 and Hepatitis C virus.135

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Given the widespread health effects of chlamydial infections122 and the failed antibiotic prophylaxis of C. pneumoniae-induced exacerbation of insulin resistance and cardiovascular disease,42,43 vaccination may be an attractive and realistic approach to control most of the Chlamydophilaassociated diseases in human beings. Such vaccination approaches may have to balance the potential for exacerbated tissue damage at the primary infection site against the benefits gained by limiting chlamydial infections and thereby minimizing whole-host negative effects. In summary, infections with Chlamydophila spp. are associated with a wide range of typically asymptomatic disease conditions in both humans and animals.24,122 One of the key advances in the understanding of chlamydial infections has been the development of molecular detection methods for these pathogens with high sensitivity and specificity.109,126,136 Research leading to effective strategies for prevention and therapy of chlamydial infections, such as development of effective vaccines, will have an enormous impact on the health of humans and animals. While tremendous progress in the understanding of basic biology and molecular make-up of chlamydiae has been made over the last two decades, this progress has not translated into clinically significant improvements in prophylaxis and therapy. The main progress in understanding both human and animal chlamydial diseases, with actionable consequences in the clinical setting, has come from improvements in diagnostics. The ideal diagnostic tool should allow a rapid diagnosis at low cost, and with high specificity and sensitivity. The main challenge for PCR-based detection methods is the efficient extraction of nucleic acids from low-copy specimens and their efficient transfer into the amplification mix, as well as sensitive detection and quantification of low target copy amplicons in a single-tube assay. Well-engineered platform technologies and advanced analytical methods will further increase the potential of molecular diagnostic techniques and make them reliable and universally applied tools for costeffective rapid detection of Chlamydiae including simultaneous differentiation and typing of these bacteria.

Molecular Detection of Human Bacterial Pathogens







REFERENCES





1. Moulder, J.W. et al., Genus I. Chlamydia Jones, Rake and Stearns 1945, 55AL. In Krieg, N.R., and Holt, J.G. (Editors), Bergy’s Manual of Systemic Bacteriology, vol. 1, pp. 729–739, Williams and Wilkins, Baltimore, 1984. 2. Stephens, R.S., Introduction. In Stephens, R.S. (Editor), Chlamydia, Intracellular Biology, Pathogenesis, and Immunity, American Society for Microbiology, Washington, DC, 1999. 3. Amann, R.N. et al., Obligate intracellular bacterial parasites of acanthamoebae related to Chlamydia spp., Appl. Environ. Microbiol., 63, 115, 1997. 4. Pace, N.R., A molecular view of microbial diversity and the biosphere, Science, 276, 734, 1997. 5. Hugenholtz, P., Goebel, B.M., and Pace, N.R., Impact of culture-independent studies on the emerging phylogenetic view of bacterial diversity, J. Bacteriol., 180, 4765, 1998.





6 Kaltenboeck, B., Kousoulas, K.G., and Storz, J., Structures of and allelic diversity and relationships among the major outer membrane protein (ompA) genes of the four chlamydial species, J. Bacteriol., 175, 487, 1993. 7 Everett, K.D.E., Bush, R.M., and Andersen, A.A., Emended description of the order Chlamydiales, proposal of Parachlamydiaceae fam. nov. and Simkaniaceae fam. nov., each containing one monotypic genus, revised taxonomy of the family Chlamydiaceae, including a new genus and five new species, and standards for the identification of organisms, Int. J. Syst. Bacteriol., 49, 415, 1999. 8. Everett, K.D., and Andersen, A.A., Identification of nine species of the Chlamydiaceae using PCR-RFLP, Int. J. Syst. Bacteriol., 49, 803, 1999. 9. Bush, R.M., and Everett, K.D.E., Molecular evolution of the Chlamydiaceae, Int. J. Syst. Evol. Microbiol., 51, 203, 2001. 10. Horn, M. et al., Neochlamydia hartmannellae gen. nov., sp. nov. (Parachlamydiaceae), an endoparasite of the amoeba Hartmannella vermiformis, Microbiology, 146, 1231, 2000. 11. Schachter, J. et al., Radical changes to chlamydial taxonomy are not necessary just yet, Int. J. Syst. Evol. Microbiol., 51, 249, 2001. 12. Stephens, R.S. et al., Divergence without difference: Phylogenetics and taxonomy of Chlamydia resolved, FEMS Immunol. Med. Microbiol., 55, 115, 2009. 13 Matsumoto, A., Electron microscopic observations of surface projections on Chlamydia psittaci reticulate bodies, J. Bacteriol., 151, 1040, 1982. 14. Grayston, J.T. et al., A new Chlamydia psittaci strain, TWAR, isolated in acute respiratory tract infections, N. Engl. J. Med., 315, 161, 1986. 15. Glassick, T., Giffard, P., and Timms P, Outer membrane protein 2 gene sequences indicate that Chlamydia pecorum and Chlamydia pneumoniae cause infections in koalas, Syst. Appl. Microbiol., 19, 457, 1996. 16. Storey, C., Evidence for Chlamydia pneumoniae of nonhuman origin, J. Gen. Microbiol., 139, 2621, 1993. 17. Berger, L. et al., Chlamydia pneumoniae in a free-ranging giant barred frog (Mixophyes iteratus) from Australia, J. Clin. Microbiol., 37, 2378, 1999. 18. Hotzel, H. et al., Genetic characterization of a Chlamydophila pneumoniae isolate from an African frog and comparison to currently accepted biovars, Syst. Appl. Microbiol., 24, 63, 2001. 19. Kuo, C.C. et al., Chlamydia pneumoniae (TWAR), Clin. Microbiol. Rev., 8, 451, 1995. 20. Boman, J., et al., High prevalence of Chlamydia pneumoniae DNA in peripheral blood mononuclear cells in patients with cardiovascular disease and in middle-aged blood donors, J. Infect. Dis., 178, 274, 1998. 21. Kleemola, M. et al., Epidemics of pneumonia caused by TWAR, a new Chlamydia organism, in military trainees in Finland, J. Infect. Dis., 157, 230, 1988. 22. Mayer, K.F., The host spectrum of psittacosis-lymphogranuloma venereum (PL) agents, Am. J. Ophthalmol., 63, 1225, 1967. 23. Kaleta, E.F., and Taday, E.M., Avian host range of Chlamydophila spp. based on isolation, antigen detection and serology, Avian Pathol., 32, 435, 2003. 24. Schachter, J., Infection and disease epidemiology. In Stephens, R.S. (Editor), Chlamydia, Intracellular Biology, Pathogenesis, and Immunity, p. 139, American Society for Microbiology, Washington, DC, 1999. 25. Mayer, K.F., and Eddie, B., Human carrier of the psittacosis virus, J. Infec. Dis., 88, 109, 1951.

Chlamydophila 26. Palmer, S.R., Psittacosis in man-recent developments in UK: A review, J. R. Soc. Med., 75, 262, 1982. 27. Gaede, W. et al., Chlamydophila psittaci infections in humans during an outbreak of psittacosis from poultry in Germany, Zoonoses Public Health, 55, 184, 2008. 28. Laroucau, K., Chlamydial infections in duck farms associated with human cases of psittacosis in France, Vet. Microbiol., 135, 82, 2009. 29. Greig, J.R., Enzootic abortion in ewes: A preliminary note, Vet. Rec., 42, 1225, 1936. 30. Entrican, G., Buxton, D., and Longbottom, D., Chlamydial infection in sheep: Immune control versus fetal pathology, J. R. Soc. Med., 94, 273, 2001. 31. Walder, G. et al., An unusual cause of sepsis during pregnancy: Recognizing infection with Chlamydophila abortus, Obstet. Gynecol., 106, 1215, 2005. 32. Hyde, S.R., and Benirschke K., Gestational psittacosis: Case report and literature review, Mod. Pathol., 10, 602, 1997. 33. Grayston, J.T., Chlamydia pneumoniae, strain TWAR pneumoniae, Annu. Rev. Med., 43, 317, 1992. 34. Gupta, S. et al., Elevated Chlamydia pneumoniae antibodies, cardiovascular events, and azithromycin in male survivors of myocardial infarction, Circulation, 96, 404, 1997. 35. Campbell, L.A. et al., Preclinical models for Chlamydia pneumoniae and cardiovascular disease: Hypercholesterolemic mice, Clin. Microbiol. Infect., 4, S23, 1998. 36. Ekesbo, R. et al., Combined seropositivity for H. pylori and C. pneumoniae is associated with age, obesity and social factors, J. Cardiovasc. Risk, 7, 191, 2000. 37. Smieja, M. et al., Association of circulating Chlamydia pneumoniae DNA with cardiovascular disease: A systematic review, BMC Infect. Dis., 2, 21, 2001. 38. Dart, A.M., Martin, J.L., and Kay, S., Association between past infection with Chlamydia pneumoniae and body mass index, low-density lipoprotein particle size and fasting insulin, Int. J. Obes. Relat. Metab. Disord., 26, 464, 2002. 39. Nabipour, I. et al., The association of metabolic syndrome and Chlamydia pneumoniae, Helicobacter pylori, cytomegalovirus, and herpes simplex virus type 1: The Persian Gulf Healthy Heart Study, Cardiovasc. Diabetol., 5, 25, 2006. 40. Fernández-Real, J.M. et al., Burden of infection and insulin resistance in healthy middle-aged men, Diabetes Care, 29, 1058, 2006. 41. Wang, C., Gao, D., and Kaltenboeck, B., Acute Chlamydia pneumoniae reinfection accelerates the development of insulin resistance and diabetes in obese C57BL/6 mice, J. Infect. Dis., 200, 279, 2009. 42. Cannon, C.P. et al., Antibiotic treatment of Chlamydia pneumoniae after acute coronary syndrome, N. Engl. J. Med., 352, 1646, 2005. 43. Grayston, J.T. et al., Azithromycin for the secondary prevention of coronary events, N. Engl. J. Med., 352, 1637, 2005. 44. Vanrompay, D., et al., Serotyping of European isolates of Chlamydia psittaci from poultry and other birds, J. Clin. Microbiol., 31, 134, 1993. 45. Andersen, A.A., Two new serovars of Chlamydia psittaci from North American birds, J. Vet. Diag. Invest., 9, 1997. 46. Sachse, K. et al., Genotyping of Chlamydophila psittaci using a new DNA microarray assay based on sequence analysis of ompA genes, BMC Microbiol., 8, 63, 2008. 47. Sachse, K et al., DNA microarray-based genotyping of Chlamydophila psittaci strains from cell culture and clinical samples, Vet. Microbiol., 135, 22, 2009.

533 48. Olson, B.J., and Treuting, W.L., An epidemic of a severe pneumoniae in the Bayou Region of Louisiana. IV. A preliminary note on etiology, Public Health Rep., 131, 376, 1944. 49. Schachter, J., and Dawson, C.R., Human Chlamydial Infections, PSG Publishing Co., Littleton, 1978. 50. Giroud, P., and Jadin, J., Premiers resultats concernant le virus des Bashi isole dans la provence du Kivu au Congo Belge, Bull. Soc. Pathol. Exot., 47, 578, 1954. 51. Mare, C.J., Mammalian chlamydiosis. In Beran, G.W. (Editor), Handbook of Zoonoses, p. 403, CRC Press, Boca Raton, 1994. 52. Jorgensen, D.M., Gestational psittacosis in a Montana sheep rancher, Emerg. Infect. Dis., 3, 191, 1997. 53. Villemonteix, P. et al., Pregnancy complicated by severe Chlamydia psittaci infection acquired from a goat flock: Case report, Eur. J. Obstet. Gynecol. Reprod. Biol., 37, 91, 1990. 54. Pospischil, A. et al., Abortion in humans caused by Chlamydophila abortus (Chlamydia psittaci serovar 1), Schweiz Arch. Tierheilkd, 144, 463, 2002. 55. Enright, J.B., and Sadler, W.W., Presence in human sera of complement-fixing antibodies to the virus of sporadic bovine encephalomyelitis, Proc. Soc. Exp. Biol. Med., 85, 466, 1954. 56. Barnes, M.G., and Brainerd, H., Pneumonitis with alveolarcapillary block in a cattle rancher exposed to epizootic bovine abortion, N. Engl. J. Med., 271, 981, 1964. 57. Page, L.A., and Smith, P.C., Placentitis and abortion in cattle inoculated with chlamydiae isolated from aborted human placental tissue, Proc. Soc. Exp. Biol. Med., 146, 269, 1974. 58. Kaltenboeck, B., Kousoulas, K.G., and Storz, J., Detection and strain differentiation of Chlamydia psittaci mediated by a two-step polymerase chain reaction, J. Clin. Microbiol., 29, 1969, 1991. 59. Fukushi, H., and Hirai, K., Proposal of Chlamydia pecorum sp. nov. for Chlamydia strains derived from ruminants, Int. J. Syst. Bacteriol., 42, 306, 1992. 60. Jackson, M. et al., Epizootiology of Chlamydia infections in two free-range koala populations, Vet. Microbiol., 65, 255, 1999. 61. Baker, J.A., A virus causing pneumonia in cats and producing elementary bodies, J. Exp. Med., 79, 159, 1944. 62. Cello, R.M., Ocular infections in animals with PLT (Bedsonia) group agents, Am. J. Opthalmol., 63, 1270, 1967. 63. Shewen, P.E., Povey, R.C., and Wilson, M.R., Feline chlamydial infection, Can. Vet. J., 19, 289, 1978. 64. Murray, E.S., Guinea pig inclusion conjunctivitis virus. I. Isolation and identification as a member of psittachosis-granuloma-trachoma group, J. Infect. Dis., 114, 1, 1964. 65. Gaede, W. et al., Detection of Chlamydophila caviae and Streptococcus equi subsp. zooepidemicus in horses with signs of rhinitis and conjunctivitis, Vet. Microbiol., 142, 440, 2010. 66. Ramsey, K.H., Alternative mechanisms of pathogenesis. In Bavoil, P.M., and Wyrick, B. (Editors), Chlamydia Genomics and Pathogenesis, p. 435, Horizon Bioscience Press, Norfolk, 2006. 67. Morrison, R.P., Feilzer, K., and Tumas, D.B., Gene knockout mice establish a primary protective role for major histocompatibility complex class II-restricted responses in Chlamydia trachomatis genital tract infection, Infect. Immun., 63, 4661, 1995. 68. Rank, R.G., The role of the CD4 T cell in the host response to Chlamydia. In Bavoil, P.M., and Wyrick, B. (Editors), Chlamydia Genomics and Pathogenesis, p. 365, Horizon Bioscience Press, Norfolk, 2006. 69. Natividad, et al., Genetic variation at the TNF locus and the risk of severe sequelae of ocular Chlamydia trachomatis infection in Gambians, Genes Immun., 8, 288, 2007.

534 70. Perry, L.L., Feilzer, K., and Caldwell, H.D., Immunity to Chlamydia trachomatis is mediated by T helper 1 cells through IFN-gamma-dependent and -independent pathways, J. Immunol., 158, 3344, 1997. 71. Rottenberg, M.E. et al., Regulation and role of IFN-gamma in the innate resistance to infection with Chlamydia pneumoniae, J. Immunol., 164, 4812, 2000. 72. Swindle, E.J., and Metcalfe, D.D., The role of reactive oxygen species and nitric oxide in mast cell-dependent inflammatory processes, Immunol. Rev., 217, 186, 2007. 73. Huang, J. et al., The quantity of nitric oxide released by macrophages regulates Chlamydia-induced disease, Proc. Natl. Acad. Sci. USA, 99, 3914, 2002. 74. Ramsey, K.H. et al., Role for inducible nitric oxide synthase in protection from chronic Chlamydia trachomatis urogenital disease in mice and its regulation by oxygen free radicals, Infect. Immun., 69, 7374, 2001. 75. Li, P. et al., Amino acids and immune function, Br. J. Nutr., 98, 237, 2007. 76. Cunningham-Rundles, S., McNeeley, D.F., and Moon, A., Mechanisms of nutrient modulation of the immune response, J. Allergy Clin. Immunol., 115, 1119, 2005. 77. Field, C.J., Johnson, I.R., and Schley, P.D., Nutrients and their role in host resistance to infection, J. Leukoc. Biol., 71, 16, 2002. 78. Wintergerst, E.S., Maggini, S., and Hornig, D.H., Contribution of selected vitamins and trace elements to immune function, Ann. Nutr. Metab., 51, 301, 2007. 79. Chan, J. et al., Effects of protein calorie malnutrition on tuberculosis in mice, Proc. Natl. Acad. Sci. USA, 93, 14857, 1996. 80. McMurray, D.N., and Bartow, R.A., Immunosuppression and alteration of resistance to pulmonary tuberculosis in guinea pigs by protein undernutrition, J. Nutr., 122, 738, 1992. 81. Ward, M.E., Mechanisms of Chlamydia-induced disease. In Stephens, R.S. (Editor), Chlamydia, Intracellular Biology, Pathogenesis, and Immunity, p. 171, American Society for Microbiology, Washington, DC, 1999. 82. Thewessen, E.A. et al., Comparison of HeLa 229 and McCoy cell cultures for detection of Chlamydia trachomatis in clinical specimens, J. Clin. Microbiol., 27, 1399, 1989. 83. Johnston, S.L., and Siegel, C., Comparison of Buffalo green monkey kidney cells and McCoy cells for the isolation of Chlamydia trachomatis in shell vial centrifugation culture, Diagn. Microbiol. Infect. Dis., 15, 355, 1992. 84. Roblin, P.M. et al., Use of HEp-2 cells for improved isolation and passage of Chlamydia pneumoniae, J. Clin. Microbiol., 30, 1968, 1992. 85. Thejls, H. et al., Expanded gold standard in the diagnosis of Chlamydia trachomatis in a low prevalence population: Diagnostic efficacy of tissue culture, direct immunofluorescence, enzyme immunoassay, PCR and serology, Genitourin. Med., 70, 300, 1994. 86. Stamp, J.T. et al., Enzootic abortion in ewes: Transmission of the disease, Vet. Rec., 62, 251, 1950. 87. Aitken, I.D., and Longbottom, D., Enzootic abortion of ewes (ovine chlamydiosis). In OIE Biological Standards Commission (Editor), Manual of Diagnostic Tests and Vaccines for Terrestrial Animals (Mammals, Birds and Bees), p. 635, Office International des Epizooties, Paris, 2004. 88. Dagnall, G.J., and Wilsmore, A.J., A simple staining method for the identification of chlamydial elementary bodies in the fetal membranes of sheep affected by ovine enzootic abortion, Vet. Microbiol., 21, 233, 1990.

Molecular Detection of Human Bacterial Pathogens 89. Finlayson, J. et al., Direct immunoperoxidase method for demonstrating Chlamydia psittaci in tissue sections, J. Clin. Pathol., 38, 712, 1985. 90. Szeredi, L. et al., Intestinal chlamydia in finishing pigs, Vet. Pathol., 33, 369, 1996. 91. Juvonen, J. et al., Detection of Chlamydia pneumoniae in human nonrheumatic stenotic aortic valves, J. Am. Coll. Cardiol., 29, 1054, 1997. 92. Buxton, D. et al., Ovine chlamydial abortion: Characterization of the inflammatory immune response in placental tissues, J. Comp. Pathol., 127, 133, 2002. 93. Hotzel, H. et al., Occurrence of Chlamydiaceae spp. in a wild boar (Sus scrofa L.) population in Thuringia (Germany), Vet. Microbiol., 103, 121, 2004. 94. Wood, M.M., and Timms, P., Comparison of nine antigen detection kits for diagnosis of urogenital infections due to Chlamydia psittaci in koalas, J. Clin. Microbiol., 30, 3200, 1992. 95. Vanrompay, D. et al., Evaluation of five immunoassays for detection of Chlamydia psittaci in cloacal and conjunctival specimens from turkeys, J. Clin. Microbiol., 32, 1470, 1994. 96. Sachse, K. et al., Detection of Chlamydia suis from clinical specimens: Comparison of PCR, antigen ELISA, and culture, J. Microbiol. Methods, 54, 233, 2003. 97. Wang, S.-P., and Grayston, J.T., Human serology in Chlamydia trachomatis infection with microimmunofluorescence, J. Infect. Dis., 130, 388, 1974. 98. Verminnen, K. et al., Evaluation of a Chlamydophila psittaci diagnostic platform for zoonotic risk assessment, J. Clin. Microbiol., 46, 281, 2008. 99. Messmer, T.O. et al., Application of a nested, multiplex PCR to psittacosis outbreaks, J. Clin. Microbiol., 35, 2043, 1997. 100. Kaltenboeck, B., Schmeer, N., and Schneider, R., Evidence of numerous omp1 alleles of porcine Chlamydia trachomatis and novel chlamydial species obtained by PCR, J. Clin. Microbiol., 35, 1835, 1997. 101. Yoshida, H. et al., Differentiation of Chlamydia species by combined use of polymerase chain reaction and restriction endonuclease analysis, Microbiol. Immunol., 42, 411, 1998. 102. Sachse, K., and Hotzel, H., Detection and differentiation of chlamydiae by nested PCR, Methods Mol. Biol., 216, 123, 2003. 103. Madico, G. et al., Touchdown enzyme time release-PCR for detection and identification of Chlamydia trachomatis, C. pneumoniae, and C. psittaci using the 16S and 16S-23S spacer rRNA genes, J. Clin. Microbiol., 38, 1085, 2000. 104. Apfalter, P., Reischl, U., and Hammerschlag, M.R., In-house nucleic acid amplification assays in research: How much quality control is needed before one can rely upon the results? J. Clin. Microbiol., 43, 5835, 2005. 105. Livak, K.J. et al., Oligonucleotides with fluorescent dyes at opposite ends provide a quenched probe system useful for detecting PCR product and nucleic acid hybridization, PCR Methods Appl., 4, 357, 1995. 106. Heid, C.A. et al., Real-time quantitative PCR, Genome Res., 6, 968, 1996. 107. Wittwer, C.T. et al., Continuous fluorescence monitoring of rapid cycle DNA amplification, Biotechniques, 22, 130, 134, 1997. 108. Ehricht, R. et al., Optimized DNA microarray assay allows detection and genotyping of single PCR-amplifiable target copies, Mol. Cell. Probes, 20, 60, 2006. 109. DeGraves, F.J., Gao, D., and Kaltenboeck, B., High-sensitivity quantitative PCR platform, Biotechniques, 34, 106, 2003.

Chlamydophila 110. Geens, T. et al., Development of a Chlamydophila psittaci species-specific and genotype-specific real-time PCR, Vet. Res., 36, 787, 2005. 111. Pantchev, A. et al., New real-time PCR tests for species-specific detection of Chlamydophila psittaci and Chlamydophila abortus from tissue samples, Vet. J., 181, 145, 2009. 112. Menard, A. et al., Development of a real-time PCR for the detection of Chlamydia psittaci, J. Med. Microbiol., 55, 471, 2006. 113. Helps, C. et al., Use of real-time quantitative PCR to detect Chlamydophila felis infection, J. Clin. Microbiol., 39, 2675, 2001. 114. Pantchev, A. et al., Detection of all Chlamydophila and Chlamydia spp. of veterinary interest using species-specific real-time PCR assays, Comp. Immunol. Microbiol. Infect. Dis., 2009 (in press). 115. DeGraves, F.J. et al., Quantitative detection of Chlamydia psittaci and C. pecorum by high-sensitivity real-time PCR reveals high prevalence of vaginal infection in cattle, J. Clin. Microbiol., 41, 1726, 2003. 116. Hoffmann, B. et al., A universal heterologous internal control system for duplex real-time RT-PCR assays used in a detection system for pestiviruses, J. Virol. Methods, 136, 200, 2006. 117. Sachse, K. et al., DNA microarray-based detection and identification of Chlamydia and Chlamydophila spp., Mol. Cell. Probes, 19, 41, 2005. 118. Borel, N. et al., Direct identification of chlamydiae from clinical samples using a DNA microarray assay—a validation study, Mol. Cell. Probes, 22, 55, 2008. 119. Reinhold, P. et al., Impact of latent infections with Chlamydophila species in young cattle, Vet. J., 175, 202, 2009. 120. Bodetti, T.J. et al., Wide range of Chlamydiales types detected in native Australian mammals, Vet. Microbiol., 96, 177, 2003. 121. Borel, N. et al., Seroprevalences for ovine enzootic abortion in Switzerland, Prevent. Vet. Med., 65, 205, 2004. 122. Campbell, L.A., and Kuo, C.C., Chlamydia pneumoniae—an infectious risk factor for atherosclerosis? Nat. Rev. Microbiol., 2, 23, 2004.

535 123. Kalman, S. et al., Comparative genomes of Chlamydia pneumoniae and C. trachomatis, Nat. Genet., 21, 385, 1999. 124. Read, T.D. et al., Genome sequence of Chlamydophila caviae (Chlamydia psittaci GPIC): Examining the role of nichespecific genes in the evolution of the Chlamydiaceae, Nucleic Acids Res., 31, 2134, 2003. 125. Thomson, N.R. et al., The Chlamydophila abortus genome sequence reveals an array of variable proteins that contribute to interspecies variation, Genome Res., 15, 629, 2005. 126. Sachse, K. et al., Recent developments in the laboratory diagnosis of chlamydial infections, Vet. Microbiol., 135, 2, 2009. 127. Verkooyen, R.P. et al., Evaluation of PCR, culture, and serology for diagnosis of Chlamydia pneumoniae respiratory infections, J. Clin. Microbiol., 36, 2301, 1998. 128. Jee, J. et al., High prevalence of natural Chlamydophila species infection in calves, J. Clin. Microbiol., 42, 5664, 2004. 129. Magnino, S. et al., Chlamydial infections in feral pigeons in Europe: Review of data and focus on public health implications, Vet. Microbiol., 135, 54, 2009. 130. Kaltenboeck, B., Hehnen, H.R., and Vaglenov, A., Bovine Chlamydophila spp. infection: Do we underestimate the impact on fertility? Vet. Res. Commun., 29, 1, 2005. 131. Biesenkamp-Uhe, C. et al., Therapeutic Chlamydophila abortus and C. pecorum vaccination transiently reduces bovine mastitis associated with Chlamydophila infection, Infect. Immun., 75, 870, 2007. 132. DeGraves, F.J. et al., Reinfection with Chlamydophila abortus by uterine and indirect cohort routes reduces fertility in cattle preexposed to Chlamydophila, Infect. Immun., 72, 2538, 2004. 133. Aydemir, S. et al., The effect of Helicobacter pylori on insulin resistance, Dig. Dis. Sci., 50, 2090, 2005. 134. Mealey, B.L., and Rose, L.F., Diabetes mellitus and inflammatory periodontal diseases, Curr. Opin. Endocrinol. Diabetes Obes., 15, 135, 2008. 135. Aboud, M. et al., Insulin resistance and HIV infection: A review, Int. J. Clin. Pract., 61, 463, 2007. 136. Tong, C.Y., and Sillis, M., Detection of Chlamydia pneumoniae and Chlamydia psittaci in sputum samples by PCR, J. Clin. Pathol., 46, 313, 1993.

Chryseobacterium, 47 Elizabethkingia, and Bergeyella Dongyou Liu CONTENTS 47.1 Introduction...................................................................................................................................................................... 537 47.1.1 Classification......................................................................................................................................................... 537 47.1.2 Clinical Features................................................................................................................................................... 538 47.1.3 Diagnosis.............................................................................................................................................................. 539 47.2 Methods............................................................................................................................................................................ 540 47.2.1 Sample Preparation............................................................................................................................................... 540 47.2.2 Detection Procedures............................................................................................................................................ 540 47.3 Conclusion........................................................................................................................................................................ 541 References.................................................................................................................................................................................. 541

47.1  INTRODUCTION 47.1.1  Classification The genera Elizabethkingia, Chryseobacterium, and Bergeyella are classified in the family Flavobacteriaceae, order Flavobacteriales, class Flavobacteria, phylum Bacteroidetes. The phylum Bacteroidetes [formerly the Cytophaga–Flavobacterium–Bacteroides (CFB) group] belonging to the rRNA superfamily V is recognized as a separate line of descent within the domain Bacteria. The family Flavobacteriaceae in the CFB group consists of about 80 genera including Elizabethkingia, Chryseobacterium, Bergeyella, and Riemerella; and the genera Bergeyella, Chryseobacterium, and Riemerella appear to form a separate branch on the basis of rRNA cistron similarity and phenotypic characteristics.1–3 Members of the Bergeyella– Chryseobacterium–Riemerella branch are characterized by their lack flagellation, low G + C content (30–38 mol%), production of menaquinones as the predominant respiratory quinones, possession of large percentages of branched-chain fatty acids, absence of carbohydrate fermentation, and similarity in hydrolytic enzyme patterns. Currently, the genus Chryseobacterium (chruseos, “golden”; bacterium, a “small rod”: Chryseobacterium, “a yellow rod”) is composed of at least 45 valid species (i.e., Chryseobacterium antarcticum, Chryseobacterium anthropi, Chryseobacterium aquaticum, Chryseobacterium aquifrigidense, Chryseobacterium arothri, Chryseobacterium balustinum, Chryseobacterium bovis, Chryseobacterium caeni, Chryseobacterium daecheongense, Chryseobacterium daeguense, Chryseobacterium defluvii, Chryseobacterium flavum,

Chryseobacterium formosense, Chryseobacterium gambr­ ini, Chryseobacterium gleum, Chryseobacterium gregarium, Chryseobacterium haifense, Chryseobacterium hispanicum, Chryseobacterium hominis, Chryseobacterium humi, Chryseobacterium hungaricum, Chryseobacterium indologenes, Chryseobacterium indoltheticum, Chryseobacterium jejuense, Chryseobacterium jeonii, Chryseobacterium joos­ tei, Chryseobacterium koreense, Chryseobacterium luteum, Chryseobacterium marinum, Chryseobacterium molle, Chryseobacterium oranimense, Chryseobacterium pallidum, Chryseobacterium palustre, Chryseobacterium piscicola, Chryseobacterium piscium, Chryseobacterium scophthalmum, Chryseobacterium shigense, Chryseobacterium soldanellicola, Chryseobacterium soli, Chryseobacterium taeanense, Chryseobacterium taichungense, Chryseobacterium taiwanense, Chryseobacterium ureilyticum, Chryseobacterium vrystaatense, and Chryseobacterium wanjuense).4–22 While most Chryseobacterium species occur in aquatic environments or food products, some are pathogenic to humans and animals. For example, Chryseobacterium gleum and C. indologenes may cause nosocomial infections in humans, particularly in neonates or immunocompromised patients,23–25 and C. balustinum and C. scophthalmum have been isolated from diseased fish.26 Until recently, C. meningosepticum (formerly Flavobac­ terium meningosepticum or CDCII-a) and C. miricola were allocated to the genus Chryseobacterium. However, comparative analysis of their 16S rRNA gene sequences in relation to Chryseobacterium species led to their transfer to the new genus Elizabethkingia (named in honor of Elizabeth O. King) as Elizabethkingia meningoseptica and E. miricola. 537

538

Elizabethkingia meningoseptica is considered the most serious human pathogen in the group and is known to cause meningitis in premature and newborn infants, and pneumonia, endocarditis, postoperative bacteremia, and meningitis in adults, usually associated with a severe underlying illness. The species has been also isolated from diseased birds, frogs, turtles, and cats, whereas E. miricola was isolated from condensation water in the space station Mir. Members of the genus Chryseobacterium are strictly aerobic, nonmotile, nonspore-forming, gram-negative rods (0.5– 0.7 × 1.1–1.3 µm) that are covered by fimbriae. On LB agar, Chryseobacterium colonies measure 1.0–1.5 mm in diameter, and are circular, convex. Some species are mucoid, while others are shiny, bright yellow–colored and smooth, with a ropy consistency. They show oxidase and catalase activities, grow at 30°C and 37°C under aerobic conditions, produce acid from glucose, maltose, and ethylene glycol, produce indole, hydrolyze esculin, starch, and gelatin, display alkaline phosphatase, trypsin (benzylarginine arylamidase), and pyrrolidonyl–aminopeptidase activities. They also produce flexirubin-type pigments (except for C. hominis) and have a DNA G + C content of about 29–39 mol%. Chryseobacterium spp. occur in a variety of environments, including soil, freshand seawater, raw milk and chicken, diseased fish, bioreactor sludge, and clinical samples.7,12–14,18,19,27 Chryseobacterium indologenes is oxidase-positive, glucosenonfermenting gram-negative rod that forms smooth, circular, yellow-pigmented colonies of 1–2 mm on 5% sheep blood agar. Chryseobacterium hominis (hominis, of human being) cells are nonmotile, gram-negative rods of 1–3 µm × 1.0–1.5 µm. They grow aerobically at 20°C, 30°C, and 37°C on standard media (e.g., tryptic soy agar and blood agar), with optimal growth at 30°C, but not on MacConkey agar, cetrimide agar, or 3% NaCl agar. Colonies are circular and mucoid; some are also sticky. Some strains exhibit a pale yellow or tan pigmentation, but do not generate flexirubin pigments. They produce acid oxidatively from glucose, maltose, and ethylene glycol, but are negative for urease, lysine decarboxylase, ornithine decarboxylase, and arginine dihydrolase activities. They also produce indole, and alkalize acetate but not citrate. In addition, they show alkaline phosphatase and trypsin (benzylarginine arylamidase) activities but do not hydrolyze esculin and gelatin.19 The genus Elizabethkingia is made up of two species: Elizabethkingia meningoseptica (formerly Chryseobacterium meningosepticum, or Flavobacterium meningosepticum or CDCII-a) and Elizabethkingia miricola (formerly Chryseobacterium miricola). Elizabethkingia cells are gram-negative, nonmotile, nonspore-forming rods (0.5 × 1.0–2.5 µm). They grow well on TSA and nutrient agar at 28°C–37°C, but not at 5°C or 42°C. Colonies are white– yellow, nonpigmented, semitranslucent, circular, and shiny, with entire edges. Being catalase-, oxidase-, phosphatase-, and β-galactosidase-positive, they hydrolyze casein, esculin, and gelatin, but not starch. They do not utilize malonate, nor reduce nitrate. They produce acid from d-fructose, d-glucose,

Molecular Detection of Human Bacterial Pathogens

lactose, d-maltose, d-mannitol, and trehalose, but not from l-arabinose, d-cellobiose, raffinose, sucrose, salicin, or d-xylose. Menaquinone MK-6 is the predominant quinone. The G + C content of Elizabethkingia DNA is 35.0–38.2 mol%. Elizabethkingia meningoseptica (meninx, meningos, meninges, the membrane covering the brain; septikos putrefactive; meningoseptica, referring to association of the bacterium with both meningitis and septicemia, but not septic meningitis) was previously known as Flavobacterium meningosepticum (King 1959). E. meningoseptica cells are gramnegative, nonmotile, nonspore-forming, slender, slightly curved rods (0.5 × 1.0–2.0 µm). They show catalase, oxidase, and indole activities, but do not hydrolyze urea. The G + C content of E. meningoseptica DNA is 37.2 ± 0.6 mol% (37.1 mol% for the type strain). Elizabethkingia miricola (mirum, mir, peace, the name of Russian space station; -cola from incola, inhabitant; miricola, inhabitant of the Mir space station) was previously known as Chryseobacterium miricola (Li et al. 2004). E. miricola cells are gram-negative, nonmotile, nonspore-forming rods (0.5 × 1.0–2.5 µm). They grow well on MacConkey agar, producing very sticky colonies on solid medium. They hydrolyze urea. The G + C content of E. miricola DNA is 35.3 ± 0.3 mol% (35.0 mol% for the type strain). The genus Bergeyella consists of a single species Bergeyella zoohelcum (formerly Weeksella zoohelcum). B.  zoohelcum is a nonfermentative, nonspore-forming nonmotile, aerobic, gram-negative rod that grows well on blood agar but not on MacConkey’s agar. Colonies are circular, shiny, and smooth with entire edges. Biochemically, the bacterium is catalase, oxidase, and indole positive, and is nonsaccharolytic. B. zoohelcum is susceptible to penicillin, a feature that aids its differentiation from other similar bacteria (e.g., Flavobacterium and Sphingobacterium spp.). It is found as part of the normal oral flora of dogs, cats, and some other animals. Most clinical isolates come from infected animal bite wounds.5,28–32

47.1.2  Clinical Features Elizabethkingia meningoseptica is a gram-negative rod that is widely distributed in the environment. As a former member of the genus Chryseobacterium (Chryseobacterium meningosepticum), E. meningoseptica is known to cause meningitis and bacteremia, predominantly in premature newborns and infants. Among neonates, meningitis predominates the cases of infection with a 57% mortality rate, followed by sepsis and pneumonia.33 Among the postneonatal group, pneumonia is the most frequent infection, followed by sepsis, meningitis, endocarditis, cellulitis, abdominal infections, eye infections, sinusitis, bronchitis, and epididymitis.24,34–39 E. meningoseptica outbreaks in neonatal intensive care and hemodialysis units and its transmission in long-term acute care hospitals with mechanically ventilated patients have been reported.40–42 Occasionally, E. meningoseptica is responsible for bacterial meningitis in adults and adolescents, and cellulitis in immunocompromised patients.39,43 Being resistant to multiple

539

Elizabethkingia, Chryseobacterium, and Bergeyella

antibiotics that are prescribed for treating gram-negative bacterial infections, including extended-spectrum β-lactam agents and aminoglycosides, the bacterium represents a significant clinical concern. Ozkalay et al.43 reported a case of community-acquired E. meningoseptica meningitis and sepsis in a 17-year-old boy with thalassemia major who underwent splenectomy. E. meningosepticum was isolated in the cerebrospinal fluid (CSF) and blood samples of this patient, who recovered completely after receiving vancomycin therapy for 21 days. Bomb et al.38 documented an unusual case of native valve endocarditis in a 58-year-old man due to E. meningoseptica. Tuon et al.39 described a case of E. meningoseptica cellulitis with severe sepsis and hepatitis involving a 36-year-old male patient. The patient had a 7-day history of myalgia, fever, nausea, and vomiting before admission to hospital, and presented a right-shoulder cellulitis, tachycardia, tachypnoea, and hepatomegaly on physical examination. Laboratory tests revealed leukocytosis, renal failure, hepatitis, and metabolic acidosis. An E. meningoseptica strain was later identified in blood sample by a semiautomated system – Walkway (DadeBehring). The patient recovered after undergoing ciprofloxacin treatment. Chryseobacterium spp. are gram-negative, aerobic, motile rods that exist in soil, plants, foodstuffs, and water sources. As an opportunistic human pathogen, C. indologenes can cause bacteremia, wound sepsis, and ventilator-associated pneumonia in immunocompromised patients and patients that have received long-term broad-spectrum antibiotics via contaminated fluid-containing apparatuses and indwelling devices.44,45 Cascio et al.46 reported a C. indologenes bacteremia in a 2-year-old boy with type 1 diabetes mellitus. Two blood cultures on a standard blood culturing system (BACTEC 9120; Becton Dickinson) yielded C. indologenes isolates at day 5 after inoculation. Both isolates failed to produce acid from d-xylose and l-arabinose and were negative for esculin hydrolysis within 4 h of incubation but positive after 24 h of incubation. The biochemical profiles produced by the API 20NE system (BioMérieux) and Vitek GNI card using the Vitek-2 system (BioMérieux) confirmed the organism as C. indologenes. After treatment with ceftriaxone, the patient became afebrile after 48 h, and his general condition improved within 36 h. Bergeyella zoohelcum is commonly isolated from the upper respiratory tract of dogs, cats, and other mammals. In one study, the organism was identified in 38%–90% of nasal and oral fluids and gingival scrapings of dogs. The bacterium is an occasional zoonotic pathogen typically associated with animal bites, and is known to cause cellulitis, leg abscess, tenosynovitis, septicemia, pneumonia, and meningitis in humans.47 Shukla et al.48 reported the isolation and characterization of a fastidious Bergeyella zoohelcum strain from acute cellulitis in the upper extremity of a 60-year-old woman. The isolate was characterized by PCR amplification and sequencing of the 16S rRNA gene with broad-range eubacterial primers. Recently, Lin et al.49 described a 73-year-old man with liver cirrhosis who had no history of dog bite but had dog exposure. The

patient developed cellulitis of the left lower leg, and B. zoohelcum was isolated from blood culture. This patient was treated with cefazolin and gentamicin with a good outcome.

47.1.3  Diagnosis Phenotypic Identification. Members of the genera Elizabethkingia, Chryseobacterium, and Bergeyella are cultured on 5% sheep blood, eosin-methylene blue, and chocolate agars at 35°C in 5%–10% CO2 for 24–48 h. Initial identification is based on Gram stain, and cell and colony morphology.3 Specifically, cell morphologies are observed under a light microscope (1000× magnification) with cells grown in Anacker and Ordal’s broth for 1–3 days. Gram reaction may be based on a nonstaining method (3% KOH).50,51 Motility is examined by phase-contrast microscopy. Cellular morpho­ logy may be observed by transmission electron microscopy (TEM) with bacterial cells grown for 18 h on LB agar. The presence of flexirubin-type pigments is tested by observing the color shift when a small mass of bacterial cells collected on agar and deposited on a glass slide is flooded with a 20% (w/v) KOH aqueous solution.19 Hemolysis is recorded on trypticase soy agar supplemented with 5% (v/v) sheep blood (bioMérieux). Trypticase soy agar is also used to test the temperature tolerance at 5°C, 26°C, 37°C, and 42°C. Bacterial growth is tested on MacConkey no. 3 (Oxoid), thiosulfate-citrate-bile-sucrose (TCBS; Difco), brain heart infusion (Difco), Mueller Hinton (Difco), marine 2216 (Difco), and cetrimide. Oxidase activity is tested on oxidase test discs (bioMérieux Bactident) or oxidase test strips (Merck); and catalase activity, with 3% H2O2. The type of respiratory metabolism is determined by inoculating deep tubes of meat-liver medium containing 6% agar (BIO-RAD) Acid production is assessed using 1% glucose, H2S production using peptone iron agar (Difco 289100) and starch hydrolysis using starch agar (Difco 272100). Hydrolysis of sodium alginate, deoxyribonucleic acid, gelatin, and Tween 80 are tested both on plain nutrient agar (containing 0.5% NaCl) and on the same medium enriched to 1.5% NaCl; production of indole and reduction of nitrate are also evaluated on nutrient agar. Further biochemical identification is performed by using API 20NE and API ID32 GN microtest systems (bioMerieux). Susceptibilities to antimicrobial agents are determined by E-test strips (AB Biodisk).19,51–53 Infections are deemed community acquired if the patient admitted with an acute illness and initial cultures at the time of presentation yield a positive result. Infections are considered nosocomial if cultures are negative at the time of admission or symptomatic infections develop after the first 72 h of hospitalization. Genotypic Identification. DNA–DNA hybridization and 16S rRNA gene sequence analysis have proven useful for identification of members of the genera Elizabethkingia, Chryseobacterium, and Bergeyella.54,55 Broad-range eubacterial primers (e.g., FD1: 5′-AGA GTT TGA TCC TGG CTC AG-3′ and RD1: 5′-AAG GAG GTG ATC CAG CC-3′) are used to amplify an ~1500-bp product from the 16S rRNA

540

gene. The resulting PCR fragment is column purified and then sequenced from both sense and antisense directions by cycle sequencing.48 For genotyping purpose, pulsed-field gel electrophoresis (PFGE) and infrequent-restriction-site PCR (IRS-PCR) may be performed. PFGE and IRS-PCR demonstrate a similar efficiency in the discrimination of Elizabethkingia meningoseptica, leading to the identification of 11 distinct genotypes among the 11 clinical isolates in one study.49 Ribotyping is also useful for epidemiological studies concerning E. meningoseptica.56

47.2  METHODS 47.2.1  Sample Preparation Blood and CSF specimens are cultured on 5% sheep blood, eosin-methylene blue and chocolate agars at 35°C in 5%–10% CO2 for 24–48 h. Cell and colony morphology and biochemical properties are examined using phenotypical procedures.43 The template DNA for 16S rRNA gene analysis may be prepared by boiling a few isolated colonies in 200 μL of sterile water for 10 min. The sample is centrifuged at 15,300 × g for 2 min to pellet the cell debris. The supernatant is collected and used directly as the template for PCR amplification. Alternatively, DNA is extracted with the DNeasy tissue kit (Qiagen).

47.2.2  Detection Procedures (i) 16S rRNA Gene Analysis Protocol of Kim et al.12  Kim et al.12 utilized primers 9F (5′-GAGTTTGATCCTGGCT CAG-3′; positions 9–27 on Escherichia coli 16S rRNA numbering) and 1512R (5′-ACGGCTACCTTGTTACGA CTT-3′; positions 1512–1492) to amplify the complete 16S rRNA gene. They then used primers 9F (5′-GA GTTTGATCCTGGCTCAG-3′; 9–27), 341F (5′-CCTA CGGGAGGCAGCAG-3′; positions 341–357), 519F (5′-CAGCAGCCGCGGTAATAC-3′; positions 519–536), 907F (5′-AAACTCAAAKGAATTGACGG-3′; positions 907–926), 536R (5′-GTATTACCGCGGCTGCTG-3′; positions 536–519), 1100R (5′-GGGTTGCGCTCGTTG-3′; positions 1114–1100), and 1512R (5′-ACGGCTACCTT GTTACGACTT-3′; positions 1512–1492) to analyze the nucleotide sequence. Procedure 1. PCR mixture (100 µL) consists of 1 µM each of primers 9F and 1512R, 0.1 µM each dNTP, 10× reaction buffer, 2.5 U Taq DNA polymerase, and 100 ng extracted DNA. 2. PCR amplification is conducted with 35 cycles of 94°C for 1 min, 60°C for 1 min, and 72°C for 2 min, followed by a final extension at 72°C for 10 min. 3. The amplified fragment is then sequenced using primers 9F, 519F, 907F, 536R, 1100R, and 1512R.

Molecular Detection of Human Bacterial Pathogens



4. The full 16S rRNA gene sequences are compiled using SeqMan software and sequences of the test strains are edited using the BioEdit program and aligned using CLUSTAL X. The distance matrix is calculated by the BioEdit program after deleting regions containing ambiguous nucleotides. The phylogenetic tree is constructed by the neighbor-joining method using the Mega2 program.

(ii) 16S rRNA Gene Analysis Protocol of Han et al.32  Han et al.32 employed universal 16S rRNA gene primers A17F and 1512R for PCR amplification followed by DNA sequencing to identify Elizabethkingia, Chryseobacterium, and Bergeyella bacteria (Table 47.1). Procedure 1. PCR mixture (25-μL) is made up of GoTaq DNA polymerase (Promega), and 2 μL of DNA. Amplification is performed with an initial denaturation at 94°C for 3 min; 28–32 cycles of 94°C for 1 min, 50°C for 1 min, and 72°C for 2 min; and a final extension at 72°C for 10 min, in an Applied Biosystems 2720 thermal cycler. 2. The PCR products are visualized by 1% agarose gel electrophoresis. The PCR products are then cloned into the pCR11 vector, and a total of 10 clones with positive 16S rRNA gene inserts are selected for species identification. 3. Plasmids from these 10 clones are extracted by using a WizardPlus SV Minipreps DNA purification system (Promega), and their inserts are sequenced by using primers GW1 and GW. 4. The sequences are assembled and aligned by using the VectorNTI program (Invitrogen). The NCBI BLAST nucleotide sequence database is searched for preliminary species identification. The 16S rRNA gene sequences are aligned by using the ClustalX1.8 program. A tree for phylogenetic analysis is constructed by using Molecular Evolutionary Genetics Analysis (version 4) software by calculation of a distance matrix and tree reconstruction by the neighbor-joining method. The statistical robustness of the analysis is estimated by bootstrapping with 1000 replicates.

TABLE 47.1 Primers for Amplification and Sequencing Analysis of Elizabethkingia, Chryseobacterium, and Bergeyella rRNA Gene Primer A17F 1512R GW1 GW2

Sequence (5′→3′) GTTTGATCCTGGCTCAG TACCTTGTTACGACTT GTTGCAACAAATTGATGAGCAATGC GTTGCAACAAATTGATGAGCAATTA

Purpose PCR PCR DNA sequencing DNA sequencing

541

Elizabethkingia, Chryseobacterium, and Bergeyella

Sequences are considered monophylic when they have a connecting node within a genetic distance of 0.05. This analysis allows the presumptive identification of the bacteria by inferring the evolutionary relationships of homologous sequences. The similarities between the 16S rRNA genes within a species are expected to be >97%, and those between different species within a genus are between 93.3% and 99.9%.







47.3  CONCLUSION The genera Elizabethkingia, Chryseobacterium, and Bergeyella in the family Flavobacteriaceae encompass a number of nonmotile, catalase-positive, oxidase-positive, indolepositive, nonglucose-fermenting, gram-negative bacilli that are distributed in a variety of environments. Some of these bacteria (e.g., Elizabethkingia meningoseptica, formerly Flavobacterium meningosepticum or Chryseobacterium meningosepticum; Chryseobacterium indologenes, and Bergeyella zoohelcum, formerly Weeksella zoohelcum) are associated with clinical diseases in humans, especially in immunosuppressed individuals and patients with underlying disease. The clinical symptoms of E. meningoseptica range from meningitis, bacteremia, pneumonia, endocarditis, cellulitis, abdominal infections, eye infections, sinusitis, bronchitis, and epididymitis.33–49,57 Considering that E.  meningoseptica is largely tolerant of antibiotics that are used presently to treat microbial infections (e.g., β-lactam agents and aminoglycosides), it is important to correctly and rapidly identify the bacterium for selecting appropriate antibiotic therapy. Overcoming the time consuming and variable nature of phenotypic procedure, nucleic acid–amplification technologies such as PCR provide a highly sensitive and precise alternative for laboratory identification and detection of Elizabethkingia, Chryseobacterium, and Bergeyella organisms. Further application of these state-of-art techniques will no doubt contribute to a better understanding of the roles of these bacteria in human diseases as well as to the designing of more effective control measures against these pathogens of increasing importance.







REFERENCES

1. Woese, C.R. et al., The Flexibacter-Flavobacterium connection, Syst. Appl. Microbiol., 13, 161, 1990. 2. Hugo, C.J. et al., A polyphasic taxonomic study of Chryseobacterium strains isolated from dairy sources, Syst. Appl. Microbiol., 22, 586, 1999. 3. Jooste, P.J., and Hugo, C.J., The taxonomy, ecology and cultivation of bacterial genera belonging to the family Flavobacteriaceae, Int., J., Food Microbiol., 53, 81, 1999. 4. Yabuuchi, E. et al., Sphingobacterium gen. nov., Sphingobacterium spiritivorum comb. nov., Sphingobacterium multivorum comb. nov., Sphingobacterium mizutae sp. nov., and Flavobacterium indologenes sp. nov. glucose-nonfermenting gram-negative rods in CDC groups IIK-2 and IIb, Int. J. Syst. Bacteriol., 33, 580, 1983.







5. Vandamme, P. et al., New perspectives in the classification of the flavobacteria: Description of Chryseobacterium gen. nov., Bergeyella gen. nov., and Empedobacter nom. rev., Int. J. Syst. Bacteriol., 44, 827, 1994. 6. Bernardet, J.-F. et al., Cutting a Gordian knot: emended classification and description of the genus Flavobacterium, emended description of the family Flavobacteriaceae, and proposal of Flavobacterium hydatis nom. nov. (basonym, Cytophaga aquatilis Strohl and Tait 1978), Int. J. Syst. Bacteriol., 46, 128, 1996. 7. Bernardet, J.-F., Nakagawa, Y., and Holmes, B., Proposed minimal standards for describing new taxa of the family Flavobacteriaceae and emended description of the family, Int. J. Syst. Evol. Microbiol., 52, 1049, 2002. 8. Hugo, C.J. et al., Chryseobacterium joostei sp. nov., isolated from the dairy environment, Int. J. Syst. Evol. Microbiol., 53, 771, 2003. 9. Kampfer, P. et al., Chryseobacterium defluvii sp. nov., isolated from wastewater, Int. J. Syst. Evol. Microbiol., 53, 93, 2003. 10. Kampfer, P. et al., Description of Chryseobacterium anthropi sp. nov. to accommodate clinical isolates biochemically similar to Kaistella koreensis and Chryseobacterium haifense, proposal to reclassify Kaistella koreensis as Chryseobacterium koreense comb. nov. and emended description of the genus Chryseobacterium, Int. J. Syst. Evol. Microbiol., 59, 2421, 2009. 11. Li, Y. et al., Chryseobacterium miricola sp. nov., a novel species isolated from condensation water of space station Mir, Syst. Appl. Microbiol., 26, 523, 2003. 12. Kim, M.K. et al., Kaistella koreensis gen. nov., sp. nov., a novel member of the Chryseobacterium–Bergeyella–Riemerella branch, Int. J. Syst. Evol. Microbiol., 54, 2319, 2004. 13. Kim, K.K. et al., Chryseobacterium daecheongense sp. nov., isolated from freshwater lake sediment, Int. J. Syst. Evol. Microbiol., 55, 133, 2005. 14. Kim, K.K. et al., Transfer of Chryseobacterium meningosepticum and Chryseobacterium miricola to Elizabethkingia gen. nov. as Elizabethkingia meningoseptica comb. nov. and Elizabethkingia miricola comb. nov., Int. J. Syst. Evol. Microbiol., 55, 1287, 2005. 15. de Beer, H. et al., Chryseobacterium vrystaatense sp. nov., isolated from raw chicken in a chicken-processing plant, Int. J. Syst. Evol. Microbiol., 55, 2149, 2005. 16. de Beer, H. et al., Chryseobacterium piscium sp. nov., isolated from fish of the South Atlantic Ocean off South Africa, Int. J. Syst. Evol. Microbiol., 56, 1317, 2006. 17. Shimomura, K., Kaji, S., and Hiraishi, A., Chryseobacterium shigense sp. nov., a yellow-pigmented, aerobic bacterium isolated from a lactic acid beverage, Int. J. Syst. Evol. Microbiol., 55, 1903, 2005. 18. Quan, Z.-X. et al., Chryseobacterium caeni sp. nov., isolated from bioreactor sludge, Int. J. Syst. Evol. Microbiol., 57, 141, 2007. 19. Vaneechoutte, M. et al., Chryseobacterium hominis sp. nov., to accommodate clinical isolates biochemically similar to CDC groups II-h and II-c, Int. J. Syst. Evol. Microbiol., 57, 2623, 2007. 20. Hantsis-Zacharov, E. et al., Chryseobacterium oranimense sp. nov., a psychrotolerant, proteolytic and lipolytic bacterium isolated from raw cow’s milk, Int. J. Syst. Evol. Microbiol., 58, 2635, 2008. 21. Yassin, A.F. et al., Chryseobacterium treverense sp. nov., isolated from human clinical source, Int. J. Syst. Evol. Microbiol., 60, 1993, 2010.

542 22 Pires, C. et al., Chryseobacterium palustre sp. nov. and Chryseobacterium humi sp. nov., isolated from industrially contaminated sediments, Int. J. Syst. Evol. Microbiol., 60, 402, 2010. 23. Holmes, B. et al., Flavobacterium gleum, a new species found in human clinical specimens, Int. J. Syst. Bacteriol., 34, 21, 1984. 24. Bloch, K.C., Nadarajah, R., and Jacobs, R., Chryseobacterium meningosepticum: An emerging pathogen among immunocompromised adults: Report of 6 cases and literature review, Medicine, 76, 30, 1997. 25. Al-Tatari, H., Asmar, B.I., and Ang, J.Y., Lumboperitonial shunt infection due to Chryseobacterium indologenes, Pediatr. Infect. Dis. J., 26, 657, 2007. 26. Bernardet, et al., Polyphasic study of Chryseobacterium strains isolated from diseased aquatic animals, Syst. Appl. Microbiol., 28, 640, 2005. 27. Hugo, C.J., and Jooste, P.J., Preliminary differentiation of food strains of Chryseobacterium and Empedobacter using multilocus enzyme electrophoresis, Food Microbiol., 14, 133, 1997. 28. Holmes, B. et al., Weeksella zoohelcum sp. nov. (formerly group IIj) from human clinical specimens, Syst. Appl. Microbiol., 8, 191, 1986. 29. Kivinen, P.K. et al., Bergeyella zoohelcum septicaemia of a patient suffering from severe skin infection, Acta Derm. Venereol., 83, 74, 2003. 30. Beltran, A. et al., A case of Bergeyella zoohelcum bacteremia after ingestion of a dish prepared with goat blood, Clin. Infect. Dis., 42, 891, 2006. 31. Han, Y.W. et al., Transmission of an uncultivated Bergeyella strain from the oral cavity to amniotic fluid in a case of preterm birth, J. Clin. Microbiol., 44, 1475, 2006. 32. Han, Y.W. et al., Uncultivated bacteria as etiologic agents of intra-amniotic inflammation leading to preterm birth, J. Clin. Microbiol., 47, 38, 2009. 33. Güngör, S. et al., A Chryseobacterium meningosepticum outbreak in a neonatal ward, Infect. Control Hosp. Epidemiol., 24, 613, 2003. 34. Cone, L.A. et al., Osteomyelitis due to Chryseobacterium (Flavobacterium) meningosepticum, Antimicrob. Infect. Dis. Newsl., 17, 60, 1998. 35. Chiu, C.H. et al., Atypical Chryseobacterium meningosepticum and meningitis and sepsis in newborns and the immunocompromised, Taiwan, Emerg. Infect. Dis., 6, 481, 2000. 36. Lee, C.C. et al., Fatal case of community-acquired bacteremia and necrotizing fasciitis caused by Chryseobacterium meningosepticum: Case report and review of the literature, J. Clin. Microbiol., 44, 1181, 2006. 37. Lin, P.Y. et al., Clinical and microbiological analysis of bloodstream infections caused by Chryseobacterium meningosepticum in nonneonatal patients, J. Clin. Microbiol., 42, 3353, 2004. 38. Bomb, K., Arora, A., and Trehan, N., Endocarditis due to Chryseobacterium meningosepticum,Indian J. Med. Microbiol., 25, 161, 2007. 39. Tuon, F.F. et al., Chryseobacterium meningosepticum as a cause of cellulitis and sepsis in an immunocompetent patient, J. Med. Microbiol., 56, 1116, 2007.

Molecular Detection of Human Bacterial Pathogens 40. Sztajnbok, J., and Troster, E.J., Community-acquired Chryseobacterium meningosepticum pneumonia and sepsis in a previously healthy child, J. Infect., 37, 310, 1998. 41. Lim, L.C., Low, J.A., and Chan, K.M., Chryseobacterium meningosepticum (Flavobacterium meningosepticum)— a report of five cases in a local hospital, Ann. Acad. Med. Singapore, 28, 858, 1999. 42. Weaver, K.N. et al., Acute emergence of Elizabethkingia meningoseptica infection among mechanically ventilated patients in a long-term acute care facility, Infect. Control Hosp. Epidemiol., 31, 54, 2010. 43. Ozkalay, N. et al., Community-acquired meningitis and sepsis caused by Chryseobacterium meningosepticum in a patient diagnosed with thalassemia major, J. Clin. Microbiol., 44, 3037, 2006. 44. Christakis, G.B. et al., Chryseobacterium indologenes noncatheter-related bacteremia in a patient with a solid tumor, J. Clin. Microbiol., 43, 2021, 2005. 45. Douvoyiannis, M. et al., Chryseobacterium indologenes bacteremia in an infant, Int. J. Infect. Dis., 2009 September 1. 46. Cascio, A. et al., Chryseobacterium indologenes bacteraemia in a diabetic child, J. Med. Microbiol., 54, 677, 2005. 47. Montejo, M. et al., Bergeyella zoohelcum bacteremia after a dog bite, Clin. Infect. Dis., 33, 1608, 2001. 48. Shukla, S.K. et al., Isolation of a fastidious Bergeyella species associated with cellulitis after a cat bite and a phylogenetic comparison with Bergeyella zoohelcum strains, J. Clin. Microbiol., 42, 290, 2004. 49. Lin, W.R., Chen, Y.S., and Liu, Y.C., Cellulitis and bacteremia caused by Bergeyella zoohelcum, J. Formos. Med. Assoc., 106, 573, 2007. 50. Buck, J.D., Nonstaining (KOH) method for determination of Gram reactions of marine bacteria, Appl. Environ. Microbiol., 44, 992, 1982. 51. Dees, S.B. et al., Chemical and phenotypic characteristics of Flavobacterium thalpophilum compared with those of other Flavobacterium and Sphingobacterium species, Int. J. Syst. Bacteriol., 35, 16, 1985. 52. Fraser, S.L., and Jorgensen, J.H., Reappraisal of antimicrobial susceptibilities of Chryseobacterium and Flavobacterium species and method for reliable susceptibility testing, Antimicrob. Agents Chemother., 41, 2738, 1997. 53. Vessillier, S. et al., Overproduction and biochemical characterization of the Chryseobacterium meningosepticum BlaB metallo-β-lactamase, Antimicrob. Agents Chemother., 46,1921, 2002. 54. Ursing, J., and Bruun, B., Genetic heterogeneity of Flavobacterium meningosepticum demonstrated by DNADNA hybridization, Acta Pathol. Microbiol. Immunol. Scand. Sect. B, 95:33, 1987. 55. Drancourt, M., Berger, P., and Raoult, D., Systematic 16S rRNA gene sequencing of atypical clinical isolates identified 27 new bacterial species associated with humans, J. Clin. Microbiol., 42, 2197, 2004. 56. Colding, H. et al., Ribotyping for differentiating Flavobacterium meningosepticum isolates from clinical and environmental sources, J. Clin. Microbiol., 32, 501, 1994. 57. Lu, C.H. et al., An adult case of Chryseobacterium meningosepticum meningitis, Jpn. J. Infect. Dis., 57, 214, 2004.

48 Fusobacterium Dongyou Liu and Xiaoming Dong CONTENTS 48.1 Introduction...................................................................................................................................................................... 543 48.1.1 Classification, Morphology, and Biology............................................................................................................. 543 48.1.2 Clinical Features and Pathogenesis...................................................................................................................... 544 48.1.3 Diagnosis.............................................................................................................................................................. 545 48.2 Methods............................................................................................................................................................................ 546 48.2.1 Sample Preparation............................................................................................................................................... 546 48.2.2 Detection Procedures............................................................................................................................................ 546 48.2.2.1 PCR Identification of F. necrophorum.................................................................................................. 546 48.2.2.2 PCR Identification of F. nucleatum....................................................................................................... 548 48.2.2.3 PCR Identification of Fusobacterium spp............................................................................................. 549 48.3 Conclusion.........................................................................................................................................................................551 References...................................................................................................................................................................................551

48.1  INTRODUCTION 48.1.1  Classification, Morphology, and Biology The genus Fusobacterium covers a large number of grampositive, nonspore-forming, nonmotile, anaerobic, rodshaped bacterial species that come under the family Fusobacteriaceae, class Fusobacteria, phylum Fusobacteria. Currently, at least 25 species/subspecies are recognized within the genus: Fusobacterium alocis, Fusobacterium canifelinum, Fusobacterium equinum, Fusobacterium gonidiaformans, Fusobacterium mortiferum, Fusobacterium naviforme, Fusobacterium necrogenes, Fusobacterium necrophorum subsp. funduliforme, Fusobacterium necrophorum subsp. necrophorum, Fusobacterium nucleatum subsp. animalis, Fusobacterium nucleatum subsp. fusiforme, Fusobacterium nucleatum subsp. nucleatum, Fusobacterium nucleatum subsp. polymorphum, Fusobacterium nucleatum subsp. vincentii, Fusobacterium perfoetens, Fusobacterium periodonticum, Fusobacterium plautii, Fusobacterium polysaccharolyticum, Fusobacterium prausnitzii, Fusobacterium pseudonecrophorum, Fusobacterium russii, Fusobacterium simiae, Fusobacterium sulci, Fusobacterium ulcerans, and Fusobacterium varium. While members of the genus Fusobacterium form part of commensal microflora on the mucosal surfaces (e.g., mouth, upper respiratory tract, gastrointestinal tract, female genital tract) of humans and animals,1–4 some Fusobacterium spp. (e.g., F. necrophorum, F. nucleatum, F. mortiferum, and F. varium) have the capacity to invade host tissues, especially after accidental trauma, surgery, and edema, causing various

purulent or gangrenous infections in man, especially young and elderly patients.5–12 In this chapter, we focus on two most important and best characterized species of the genus Fusobacterium, that is, F. necrophorum and F. nucleatum. Fusobacterium necrophorum (fusus, a spindle; fusobacterium, spindle-shaped bacterium) cells are gram-negative, obligately anaerobic, nonmotile, beta-hemolytic, nonsporeforming, rods with rounded ends. F. necrophorum cells are often arranged singly or in pairs. Besides production of gas and indole, they generate butyric acid from peptone and glucose and propionic acid from lactate and threonine. However, they are unable to hydrolyze esculin and starch, reduce nitrate, produce acid from carbohydrates, and grow in 20% bile, in addition to being catralase-negative. Being a pleomorphic species, F. necrophorum is distinguished into two subspecies: F. necrophorum subsp. necrophorum (biovar A) and F. necrophorum subsp. funduliforme (biovar B).13 Whereas the necrophorum subspecies is associated mostly with infections in domestic mammals (e.g., calf diphtheria, labial necrosis of rabbits, necrotic rhinitis of pigs, and foot rot of cattle, sheep, and goats), the funduliforme subspecies infects mainly humans, causing Lemierre’s syndrome, a rare and once life-threatening disease that is characterized by acute pharyngitis, thrombophlebitis, and abscessation of the internal jugular vein. In addition, the funduliforme subspecies is also responsible for peritonsillar abscess and tonsillitis. Fusobacterium necrophorum subsp. necrophorum cells measure 0.5–0.6 µm by 3–70 µm. Colonies on GAM agar supplemented with 5% defibrinated horse blood are about 2  mm in diameter and appear circular, with grayish to 543

544

whitish color, and a flat and bumpy surface. The bacterium can liquefy gelatin and ferment maltose, and has a G + C content of 28–31 mol%. The type strain is strain VPI 2891 (= JCM 3718 = ATCC 25286).13 Fusobacterium necrophorum subsp. funduliforme (fundula, “sausage”; forma, “shape”: funduliforme, “sausage shaped”) cells measure 0.4–0.8 µm by 1–8 µm, with shorter cells (1–4 µm) being predominant. Colonies on GAM agar are about 1 mm in diameter, and appear circular with entire margins, grayish, translucent, and raised with smooth surfaces. The bacterium can liquefy gelatin, and ferment glucose weakly, and has a G + C content of 27–31 mol%. The type strain is strain Fn524 (= JCM 3724).13 Fusobacterium nucleatum is divided into five subspecies: nucleatum, polymorphum, fusiforme, vincentii, and animalis.14–17 Being inhabitants of the natural cavities of man and animals, F. nucleatum generates butyric acid, metabolic end products, and irritates the fibroblast of the gum, leading to necrotic lesions, abscesses, periodontal disease and bacteremia.18

48.1.2  Clinical Features and Pathogenesis A variety of clinical disease symptoms have been attributed fully or partially to F. necrophorum.20,21 These include (i) nonstreptococcal tonsillitis, particularly, recurrent sore throat. In a survey of 251 human patients with F. necrophorum infection, the most frequent site of origin was throat (179), then ear (32), and then sinusitis (8), dental (4), esophageal (1), posttraumatic/postoperative (3), and unknown (24); (ii) peritonsillar abscess, which is a precursor to Lemierre’s syndrome; (iii) deep neck space infections such as parapharyngeal or retropharyngeal abscess, in which F. necrophorum plays an accessory role; (iv) mediastinitis, which may evolve into empyema in the absence of underlying lung lesions; (v) mastoiditis, with meningitis being the most common complication accompanied by brain abscess, epidural abscess, sigmoid sinus thrombosis, subdural empyema, perisinus abscess, and transverse and cavernous sinus thrombosis; (vi) noma, which is an extreme form of necrotizing ulcerative gingivitis, with malnutrition and viral infection being the possible triggering factors; (vii) sinusitis, with some cases presenting intracranial complications such as meningitis, cavernous sinus thrombosis, and cerebral infarction; (viii) sinus thrombosis, which is a recognized complication of F. necrophorum infection, with the postanginal cases displaying internal jugular vein thrombophlebitis; (ix) cerebral abscess, meningitis, which can be aggressive, often with purulent CSF; (x) anaerobic bacteremia, which often occurs in patients with serious underlying conditions, including cancer, surgery, and chronic organ dysfunction; and (xi) septicemia, which may lead to death within 12 days in the preantibiotic era in the classic cases, or a distressing decline to death after 40–60 days in subacute cases, or a marked spike of fever following a throat infection in late metastatic manifestations, lasting weeks or months resulting in a lung abscess, septic arthritis, and liver abscess, or bacteremia without metastases.21

Molecular Detection of Human Bacterial Pathogens

Metastatic complications comprise (i) pleuropulmonary lesions such as septic pulmonary emboli, which are the most common metastatic manifestation and archetypal radiologically identifiable lesions of Lemierre’s syndrome; (ii) bone and joint manifestations such as osteomyelitis; (iii) pyomyositis and abscesses in muscle; (iv) skin and soft-tissue lesions, such as cutaneous pustules and subcutaneous abscesses; (v)  intraabdominal sepsis, such as liver abscesses; and (vi) endocarditis and pericarditis.21 Lemierre’s syndrome (or postanginal sepsis) is an exo­ genously acquired, monomicrobial, septicemic illness with metastatic abscesses secondary to a septic thrombophlebitis of the internal jugular vein infection with F. necrophorum subsp. Funduliforme.22,23 Lemierre’s syndrome is characterized by a history of recent oropharyngeal infection (acute sore throat or tonsillitis, followed by a neck mass and neck pain), clinical or radiological evidence of internal jugular vein thrombosis (predisposing to septic thrombophlebitis), a septicemia with septic emboli in lungs and other sites, and isolation of anaerobic pathogens, mainly Fusobacterium necrophorum. Lemierre’s syndrome was relatively common in the preantibiotic era, affecting previously healthy young adults (with a median age of 19 years, of which 55%–65% were male), who lacked any identified risk factor. However, it has become rare these days (affecting about 1 per million persons per year in the 1990s) due to empirical antibiotic therapy of acute pharyngitis.8,23,24 The mortality rate F. necrophorum infection ranges from 4.9% to 6.7% in patients. F. necrophorum subsp. funduliforme is likely transmitted human-to-human through close contact between carriers and susceptible individuals or enters the alimentary tract from animal sources, as the bacterium is present in the guts of herbivores such as cattle, pigs, and dogs. Among the nonspore-forming anaerobes, Fusobacterium necrophorum is unique for its strong link with clinically distinctive necrobacillosis, postanginal sepsis, or Lemierre’s syndrome. A number of virulence factors are produced by F. necrophorum to facilitate its invasion of host cells. These include: (i) leukotoxin, which is a large, secreted protein encoded by the leukotoxin operon (lktBAC) with neutrophil cytotoxicity25–33; (ii) endotoxin, which is a lipopolysaccharide known to be an important virulence factor in many gram-negative organisms, including F. necrophorum; (iii)  hemolysin, which contributes to abscess formation in F. necrophorum infection; (iv) hemagglutinin, which is one of the components of the cell surfaces of F. necrophorum subsp. necrophorum; (v) adhesins, which allow F. necrophorum attachment to host cells; (vi) platelet aggregation, which is used by F. necrophorum subsp. necrophorum to enhance its pathogenicity; (vii) synergy, in which F. necrophorum is protected by an anaerobic environment created by facultative bacteria (e.g., group C streptococci) that utilize oxygen and lower redox. In addition, Epstein–Barr virus induces immunosuppression, with a transient decrease in T-cell-mediated immunity that may predispose to bacterial infection such as F. necrophorum.21

Fusobacterium

Besides the roles of F. necrophorum virulence factors in the disease process, several external factors (e.g., host age and immune status) may also exert influence on the disease outcome. For example, F. necrophorum is known to target young adults (late teens and early 20s), producing not only Lemierre’s syndrome but also tonsillitis and peritonsillar abscess. The absence of immunity, concomitant viral infection (e.g., EBV) and genetic abnormality (e.g., single-nucleotide polymorphisms in Toll-like receptors that determine the innate immune response to bacteria) may also predispose the host to thrombophilia, then tonsillar and internal jugular vein thrombophlebitis, metastatic lesions, and severe sepsis. F. nucleatum often aggregates with most oral bacteria and plays a critical part in the formation of dental plaque as a bridge between early colonizer (gram-positive bacteria) and late colonizer (gram-negative bacteria).34–42 By scavenging oxygen and oxidative free radicals from dental plaque, F. nucleatum helps to maintain and support the conditions for major anaerobic periodontal pathogens.43,44 In addition, F. nucleatum is capable of invading the mucosal keratinocytes and inducing proinflammatory cytokines and elastase.45,46

48.1.3  Diagnosis Clinical Diagnosis. The occurrence of chills/rigors at around 4–5 days after the onset of the sore throat is a feature of Lemierre’s syndrome or necrobacillosis, indicating that the organisms are gaining access to the circulation and in the process of initiating septicemia. Other symptoms include neck pain and sometimes stiffness, tender (normally unilateral) swelling at the angle of the jaw and the sternomastoid muscle (the cord sign), reflecting the development of internal jugular venous thrombophlebitis. The markedly raised level of C-reactive protein present in Lemierre’s syndrome helps eliminate uncomplicated viral pharyngitis (e.g., EBV) as a possible diagnosis. Lemierre’s syndrome is distinguished from acute bacterial pneumonia (e.g., Staphylococcus aureus), Legionnaires’ disease, or aspiration pneumonia by the fact that lung involvement in Lemierre’s syndrome is preceded by sore throat and may be accompanied by internal jugular vein thrombosis and that initially lung lesions consist of multiple diffusely disseminated nodular lesions which rapidly cavitate. Cavitation, septic pulmonary emboli, and jugular venous thrombosis are visible on chest X-ray, CT scanning or ultrasound scanning.21 F. necrophorum may also cause false-positive results in Mycoplasma pneumoniae 16S rRNA PCR tests.47 Phenotypic Identification. F. necrophorum has a characteristic pleomorphic morphology, with filaments, short rods, and coccoid elements, which are clearly different from F. nucleatum and other gram-negative bacteria. F. necrophorum displays varied morphological features in tissue and pus (predominantly coccobacillary and often showing bipolar staining) in comparison with those in culture (polymorphic with irregularly stained rods, filamentous forms, and vesicular forms).48,49

545

For isolation of F. necrophorum from cerebral abscess pus, sample is inoculated immediately into semisolid medium that has been gassed out. Blood agar media supplemented with vitamin K, hemin, menadione, and a reducing agent such as cysteine hydrochloride provide more suitable media for cultivating Fusobacterium spp. Fastidious anaerobe agar produces the best growth of all media for Fusobacterium spp. On a suitable medium, F. necrophorum forms characteristic colonies of cream-yellow in color, and smooth, round, and entire in shape, with an odor redolent of cabbages. On horse blood agar most colonies show a narrow zone of complete hemolysis surrounding the colonies. Under long-wave UV light, colonies on fastidious anaerobe agar fluoresce with a vivid greenish yellow color.21 Biochemically, F. necrophorum produces indole, which is detectable directly from colonies on an agar plate by using the spot indole reagent p-dimethylcinnamaldehyde. In addition, F. necrophorum produces lipase on egg yolk agar. Use of commercial kits allows convenient biochemical characterization of Fusobacterium spp. (on the basis of indolepositive and alkaline phosphatase–negative activity). These include API rapid ID 32A (BioMérieux), AN-IDENT identification system (BioMérieux), Vitek ANI card (BioMérieux), and RapID-ANA II (Innovative Diagnostic Systems, Inc., Atlanta, GA). Analysis for the presence of volatile fatty acid end products by gas-liquid chromatography is also valuable. Fusobacterium spp. show a volatile fatty acid profile containing a single major peak of butyric acid, and F. necrophorum is the only Fusobacterium sp. that ferments lactate to propionate.21 Molecular Identification. Given the time consuming nature of culture-based testing for Fusobacterium spp., molecular methods have been increasingly utilized for species- and subspecies-specific determination.47,50–59 The target genes often include 16S rRNA,60–63 rpoB gene (RNA polymerase β-subunit),64,65 hemagglutinin gene,64 and the leukotoxin operon promoter-containing intergenic region.32 While the rpoB gene is specific for F. necrophorum,64 hemagglutinin gene is absent in F. necrophorum subsp. Funduliforme.54 Furthermore, F. necrophorum subsp. necrophorum and F. necrophorum subsp. funduliforme can be differentiated by 16S-23S rRNA intergenic spacer region sequence analysis, random amplified polymorphic DNA–PCR, and ribotyping analysis.66–69 Aliyu et al. described a real-time PCR targeting the rpoB gene (RNA polymerase β-subunit) sequence for detection of F. necrophorum in throat swabs of patients with sore throat. D’Ercole et al.70 showed that F. nucleatum is most frequently detected in oral specimens by using a multiplex PCR with primers from 16S rRNA gene. Shin et al.71 deve­ loped strain-specific PCR for F. nucleatum subsp. fusiforme ATCC 51190(T) and F. nucleatum subsp. vincentii ATCC 49256(T), with two pairs of primers Fs17-F14/Fs17-R14 and pFv35-F1/Fv35-R1. More recently, real-time PCR has been also employed for improved determination and quantitation of Fusobacterium spp.56,72

546

Molecular Detection of Human Bacterial Pathogens

(Nanodrop Technologies). Fusobacterium nucleic acids can be also prepared directly from clinical samples by using commercial extraction kits.

48.2  METHODS 48.2.1  Sample Preparation Sample from the disinfected tooth surface is collected with sterile paper points, transferred to fluid thioglycolate medium or Schaedler broth (Difco), and incubated at 37°C for 14 days in an anaerobic chamber (10% H2, 5% CO2, and 80% N2). The cut files and paper points are placed in cryotubes containing 1 mL of Tris-EDTA buffer. After vigorous vortexing for 1 min, the suspension is stored at −20°C for subsequent DNA extraction. Throat swabs are transported in Stuart’s Transport Medium within 24 h of sampling. They are then inoculated on Columbia agar plates containing 5% sheep blood (Becton Dickinson) and incubated overnight at 35°C in 5% CO2. Alternatively, the swabs are inoculated on plates containing 5% horse blood (SSI) v/v and incubated at 35°C in anaerobic jars containing an atmosphere of 5% CO2, 10% H2, and 85% N2. Plates are examined after incubation for 1, 2, and 4 days. Throat swabs are stored for up to 2 days at 4°C until extraction of bacterial DNA.47 Initial identification of F. necrophorum is based on colony morphology, an odor of butyric acid, the presence of a greenish fluorescence from colonies when irradiated with UV light, and a characteristic gram-negative pleomorphic morphology under the microscope, supplemented by susceptibi­ lity to kanamycin and metronidazole on primary inoculated agar plates. Subsequent β-hemolysis on agar containing 5% horse blood, resistance to vancomycin, and susceptibility to polymyxin confirm the identification of F. necrophorum.47 For DNA extraction, Fusobacterium cells from overnight anaerobic BHI broth culture are pelleted by centrifugation at 5000 × g for 10 min at 4°C. The pelleted cells are washed and resuspended in 100 mM Tris/HCl (pH 8.0), 10 mM EDTA. The cell wall is digested with lysozyme (1 mg/mL), and the resulting lysate is treated with 1% sarkosyl followed by RNase A (20 µg/mL) and pronase (50 µg/mL). The sample is incubated on ice for 20 min and then sequentially extracted twice with phenol and chloroform. The DNA from the extract is precipitated in 0.1 volume of 3 M sodium acetate pH 5.2 and an equal volume of 2-propanol. The DNA is resuspended in 10 mM Tris/HCl pH 8.0, 1 mM EDTA. The concentration of extracted DNA is determined spectrophotometrically

48.2.2  Detection Procedures 48.2.2.1  PCR Identification of F. necrophorum (i) PCR for F. necrophorum  Bennett et al.4 reported a PCR for F. necrophorum with primers targeting the leukotoxincoding lktA gene, which is unique to F. necrophorum (including F. necrophorum subsp. necrophorum and F. necrophorum subspecies funduliforme) and not present in other Fusobacterium species/subspecies.31 The F. necrophorum-specific primers lktA-up (5′-ACAATCGGAGTAGTAGGTTC-3′) and lktA-dn (5′-ATTTGGTAACTGCCACTGC-3′) amplify a 402 bp amplicon. The PCR thermal profile consists of an initial denaturation step at 94°C for 5 min, followed by 35 cycles of 94°C for 30 s, 59°C for 30 s, and 72°C for 30 s, and a final extension at 72°C for 5 min. The PCR mixture includes 400 ng/μL bovine serum albumin (BSA) (New England Biolabs) and an extra 2.5 mM of MgCl2 to reduce effects of inhibitors in the samples. PCR products from both the lktA gene is then separated electrophoretically in 1.5% agarose gels containing 0.5 μg/mL of ethidium bromide, and visualized under UV transillumination. (ii) Real-time PCR for F. necrophorum and F. necrophorum subsp. necrophorum  Aliyu et al.64 developed a quantitative real-time PCR with primers targeting the rpoB gene for detection of F. necrophorum in throat swabs collected from patients with pharyngitis. The samples positive for F. necrophorum are further analyzed with a hemagglutininrelated protein gene-specific PCR for F. necrophorum subspecies necrophorum (Table 48.1). Procedure 1. All swabs are held in Amies transport media with charcoal, refrigerated for up to 2 days at 4°C. Bacteria are harvested from the swabs by soaking in 2 mL PBS for 10 min, vortexing for 30 s and then

TABLE 48.1 Primers and Probes for Real-Time PCR Identification of F. necrophorum and F. necrophorum Subspecies funduliforme Gene

Primer

rpoB

RPO forward RPO reverse RPO probe 1 RPO probe 2 HAEMF HAEMR

Haem

Sequence (5′→3′) TCTCTACGTATGCCTCACGGATC CGATATTCATCCGAGAGGGTCTCC GAAGACATGCCTTTCTTAGAGGAC–Fluo LCRed-640–GAACTCATTTGGACGTTGTGTTA–Pho CATTGGGTTGGATAACGACTCCTAC CAATTCTTTGTCTAAGATGGAAGCGG

Position 171–193 447–424

1786–1810 2096–2071

Specificity

F. necrophorum F. necrophorum subspecies necrophorum

Fusobacterium









547

collecting the PBS from the swab by wringing out against the wall of the container. One mL of the processed PBS is transferred to a 1.5 mL Eppendorf tube and centrifuged for 10 min at 6000 × g. The supernatant is carefully decanted and the pellet resuspended in 300 µL PBS for extraction. 2. Nucleic acid is extracted from 200 µL of the PBS supernatant with a Total Nucleic Acid Isolation Kit (Roche Molecular Biochemicals) using an automated MagNA Pure Extraction System (Roche Molecular Biochemicals). The DNA is eluted in 60 µL and stored at −20°C until use. 3. PCR mixture (20 µL) is made up of 4 µL DNA extract, 2 µL LightCycler FastStart DNA master hybridization mixture (Roche Diagnostics), 3.2 µL 25 mM MgCl2 (5 mM), 0.5 µL each of 20 pmol/ µL RPO primers, 0.2 µL each of the 10 pmol/µL PRO probes and 9.4 µL RNase free water. Positive control (F. necrophorum subsp. necrophorum ATCC 25286T DNA), a contamination control (sterile water), and extraction control (sterile swab processed as above) are included in each run. PCR quantification standards (10 −4 down to 10 −9) are also included in every run of 32 samples to facilitate quantification of the bacterial load in positive samples. 4. PCR amplification is carried out in a LightCycler as follows: one cycle of denaturation at 95°C for 10 min followed by 50 cycles of 95°C for 0 s, 60°C for 2 s, and 72°C for 15 s, with a single transition rate of 20°C per s and a single fluorescence acquisition at 60°C. On completion of amplification, a single melt cycle is produced by holding the reaction at 95°C for 0 s, then at 45°C for 30 s, followed by slow heating at a transition rate of 0.1°C per s to 85°C and continuous fluorescence acquisition. 5. For specific identification of F. necrophorum subsp. necrophorum, a real-time PCR is performed on the LightCycler using FastStart DNA Master SYBR green 1 kit. PCR mixture (20 µL) is composed of 4 µL DNA extract, 2 µL of the LightCycler SYBR green kit, 2.4 µL 25 mM MgCl2 (4 mM), 0.5 µL each of 20 pmol/µL HAEM primers, and 10.6 µL RNase free water.



6. PCR amplification is conducted in a LightCycler with one cycle of denaturation at 95°C for 10 min, 50 cycles of 95°C for 0 s, 65°C for 2 s and 72°C for 15 s with a single transition rate of 20°C per s and a single fluorescence acquisition at 72°C. On completion of amplification, a single melt cycle is produced by holding the reaction at 95°C for 0 s, then at 65°C for 30 s, followed by slow heating at a transition rate of 0.1°C per s to 95°C and continuous fluorescence acquisition.

Note: Jensen et al.47 adapted this F. necrophorum species-specific real-time PCR assay for TaqMan chemistry. While the primers RPO-forward and RPO-reverse remain the same, only one TaqMan probe (RPO probe, 5′-TTGCCGGCGGAAGACATGCCTTTCTTA-3′, positions 167–193) is needed. The probe is labeled at the 5′ end with 6-carboxyfluorescein and at the 3′ end with Black Hole Quencher 1. (iii) Real-Time PCR for F. necrophorum subsp. necrophorum and subsp. funduliforme  Jensen et al.47 described a TaqMan-based real-time quantitative PCR assay, targeting the gyrB gene, to detect and discriminate between the two subspecies of F. necrophorum in the throats of healthy volunteers, and to examine the possible role of F. necrophorum in nonstreptococcal tonsillitis (NST). A primer pair (gyrB-forward and gyrB-reverse) amplifies a 306 bp fragment from both F. necrophorum subsp. necrophorum and subsp. funduliforme subspecies, which are detected with two subspecies-specific TaqMan probes (probe 1 and probe 2). The subsp. necrophorum-specific probe 1 is labeled at the 5′ end with 6-carboxyfluorescein, and at the 3′ end with Black Hole Quencher 1, whereas the subsp. funduliforme-specific probe 2 is labeled at the 5′ end with hexachlorofluorescein and at the 3′ end with Black Hole Quencher 2 (Table 48.2). The assay is carried out in two separate PCR. Procedure 1. Bacterial DNA is isolated by vortexing the swabs in 1 mL of physiological saline for 1 min and then extracting the nucleic acid from 200 µL of saline using the Kingfisher mL magnetic particle processor (Thermo Electron Corporation) and a Biosprint 15 DNA Blood Kit (Qiagen). Nucleic

TABLE 48.2 Primers and Probes for Real-Time PCR Identification of F. necrophorum subsp. necrophorum and subsp. funduliforme Gene

Primer

Sequence (5′→3′)

Position

gyrB

gyrB-forward gyrB-reverse Probe 1

AGGATTGCATGGAGTAGGAA CCTATTTCATTTCGACAATCCA FAM-TCTACTTTGGAGGTTGGAGAAACAAC-BHQ

27–46 332–311 160–185

Probe 2

HEX-TCCGCTTTAGAGGCTGGAGAAACGAC-BHQ

160–185

Specificity F. necrophorum F. necrophorum subspecies necrophorum F. necrophorum subspecies funduliforme

548





acids are eluted in 200 µL of the elution buffer supplied with the extraction kit, and stored at 4°C until use. 2. PCR for both assays is performed in 25-µL mixtures containing 12.5 µL of 2 × Brilliant QPCR Master Mix (Stratagene), 2.5 µL of 100 nM (final concentration) TaqMan probe, 2.5 µL of 300 or 400 nM (final concentrations) forward and reverse primer, for the rpoB and gyrB assays, respectively, and 5 µL of template DNA. DNA extracted from F. necrophorum subsp. necrophorum ATCC 25286 and subsp. funduliforme ATCC 51357 is included as positive controls and sterile water as negative control in each rune. 3. Thermal cycling on an Mx3000P Real-time PCR System (Stratagene) comprises 10 min at 95°C, followed by 55 cycles of 30 s at 95°C and 60 s at 60°C for both assays. All samples are run in duplicate. Results are analyzed using the Mx3000P software package (Stratagene).

Note: The minimum detection limit of the assays for both subspecies is between 1.5 × 102 and 1.5 × 103 CFU/swab (0.75 and 7.5 CFU/reaction). 48.2.2.2  PCR Identification of F. nucleatum (i) Nested PCR for F. nucleatum  Kulekci et al.38 presented a nested PCR for detection of F. nucleatum in children with persistent middle-ear effusion (MEE) as a consequence of acute otitis media (AOM) and otitis media with effusion (OME). The first-round PCR relies on E. coli 16S ribosomal RNA (rRNA) universal primers common to all bacterial species (i.e., pHF: 5′-AAGGAGGTGATCCAGCCGCA-3′ and pAF: 5′-GAGTTTGATCATGGCTCAG-3′). MEE samples showing about 1400 bp amplicon with the universal primers are further assessed by using F. nucleatum specific primers from the 16S rRNA gene sequences (i.e., FnF: 5′-CTAAAT ACGTGCCAGCAGCC-3′, positions 487–506 and FnR: 5′-CGACCCCCAACACCTAGTAA-3′, positions 821–802). Procedure 1. The external ear canal is disinfected and desquamated by placing 70% ethanol in the ear canal for 1 min. Immediately after myringotomy, the effusion is removed from the middle-ear cavity with an electronic suction device into a sterile Eppendorf tube and stored at −20°C until use. 2. A 100 µL aliquot of sample is diluted in 100 µL of 50 mM Tris-HCL pH 8.5, 50 mM EDTA, 2.5% SDS and digested overnight at 50°C after addition of 25 µL proteinase K (20 mg/mL). Sodium perchlorate is then added to a final concentration of 1 M, and the mixture is incubated at 50°C for 1 h. After addition of 200 µL of STE (150 mM NaCl, 10 mM Tris-HCl and 1 mM EDTA pH 8.5), the DNA is extracted with 400 µL phenol-chloroform (1:1), once

Molecular Detection of Human Bacterial Pathogens







with phenol-chloroform-isoamyl alcohol (25:24:1) and precipitated with ethanol, dried under vacuum, dissolved in 100 µL of sterile distilled water and stored at −20°C. 3. The PCR mixture (50 µL) is composed of 1 μM of each primer, 1.5 mM MgCl2, 200 μM of each deoxynucleoside triphosphate, 1× Taq polymerase buffer (Boehringer Mannheim), 1.25 U of Taq polymerase, and 10 µL of template DNA. F. nucleatum ATCC 49256 is used as positive control and sterile distilled as a negative control for each reaction series. 4. PCR amplification is conducted in a DNA thermal cycler (GeneAmp 9600 thermal cycler; Perkin Elmer) with an initial denaturation at 94°C for 3 min; 36 cycles of 94°C for 45 s, 55°C for 30 s, 72°C for 45 s; and a final extension at 72°C for 7 min. 5. After amplification, 10 µL of PCR products is analyzed by electrophoresis on a 2% agarose gel, stained with 0.5 µg/mL ethidium bromide and visualized by ultraviolet light illumination.

Note: The size of the PCR products is estimated using f X174 DNA-Hae III Digest as a molecular weight marker. A band of 316 bp is expected for products from the second round PCR. (ii) Multiplex PCR for F. nucleatum subsp. fusiforme and subsp. vincentii  Shin et al.71 designed strain-specific PCR primers for F. nucleatum subsp. fusiforme ATCC 51190T and F. nucleatum subsp. vincentii ATCC 49256T from the nucleotide sequence of the Fs17 (GenBank accession no., AY185204) and Fv35 (GenBank accession no., AY171472) DNA probes, respectively (Table 48.3). Procedure 1. PCR mix (20 μL) is composed of 5 nmoles each of deoxynucleoside triphosphate, 0.8 mmoles of KCl, 0.2 mmoles of Tris-HCl pH 9.0, 0.03 mmoles of MgCl2 and 1 U of Taq DNA polymerase, 20 pmoles of each primer and bacterial genomic DNA. 2. PCR is run for 32 cycles of 94°C for 1 min, 55°C or 50°C or 30 s, and 72°C for 1 min with a final 72°C for 10 min. 3. A 2 μL aliquot of the reaction mixture is then analyzed by 1.5% agarose gel electrophoresis in a TABLE 48.3 Primers for Multiplex PCR Identification of F. nucleatum subsp. fusiforme and subsp. vincentii Primer

Sequence (5′→3′)

Fs17-F14 GAT GAG GAT GAA AAG AAA CAA AGT A Fs17-R14 CCA TTG AGA AGG GCT ATT GAC Fv35-F1 ATA ATG TGG GTG AAA TAA Fv35-R1 CCC AAG GAA AAT ACT AA

Annealing Product Temperature (bp) (°C) 393

55

208

50

Fusobacterium

Tris-acetate buffer (0.04 M Tris-acetate, 0.001 M EDTA, pH 8.0) at 100 V for 30 min. The amplification products are stained with ethidium bromide, and visualized by UV transillumination. (iii) Real-Time PCR for F. nucleatum  Periasamy and Kolenbrander18 designed a real-time PCR for quantitative detection of F. nucleatum in biofilm with primers from the 16S rRNA gene: forward primer 5′-CTTAGGA ATGAGACAGAGATG-3′ and reverse primer 5′-TGATGG TAACATACGAAAGG-3′. Procedure 1. DNA is extracted from biofilms by immersing biofilm-covered pegs in 40 μL of sterile ultrapure water plus 160 μL of 0.05 M sodium hydroxide. After incubation at 60°C for 60 min, 18.4 μL of 1 M Tris-HCl pH 7.0 is added to neutralize the pH. The extract is used as the template DNA for the qPCR analyses. The bacterial genomic DNA used for standard curves is extracted from overnight cultures of F. nucleatum ATCC 10953, with a DNA extraction kit (Qiagen). Genomic DNA is stored at −20°C. 2. Quantification of F. nucleatum in the biofilms is performed by qPCR analysis using the SYBR green dye. The qPCR mixture (25 μL) is made up of 3 μL of template, 3.5 μL of diethyl pyrocarbonate-treated ultrapure water, 12.5 μL of Power SYBR green PCR master mixture (Applied Biosystems), and 3 μL each of the forward and reverse primers (375 nM). 3. qPCR is carried out with an MX3005P thermocycler (Stratagene) using the following thermal cycle recommended for the Power SYBR green PCR master mixture: 95°C for 10 min and then 40 cycles of 30 s at 95°C and 1 min at 56°C. Dissociation curves are generated by incubating reaction products at 95°C for 1 min and at 56°C for 30 s and then incrementally increasing the temperature to 95°C. Fluorescence data are collected at the end of the 56°C primer­annealing step for 40 amplification cycles and throughout the dissociation curve analysis. 4. Analysis of the melting curves with both primer sets reveals a single sharp peak. DNA concentrations (ng/mL) are calculated based on standard curves obtained by using tenfold serial dilutions of bacterial DNA isolated with a DNA extraction kit (Qiagen) and quantified with the PicoGreen fluorescence assay (Invitrogen). To convert ng DNA to numbers of cells, the following weights and genome sizes are used: 2.41 fg/genome and 2.4 Mb for fusobacteria for F. nucleatum. 48.2.2.3  PCR Identification of Fusobacterium spp. (i) Real-Time PCR for Fusobacterium spp.  Boutaga et al.72 developed a real-time PCR with the 16S rRNA gene primers and probe for detection of Fusobacterium

549

spp. in subgingival plaque samples from adult patients with periodontitis (Table 48.4). The probe is labeled with fluorescent dyes 6-carboxyfluorescein (FAM) at the 5′ end and 6-carboxytetramethylrhodamine (TAMRA) at the 3′ end. Procedure 1. PCR mixture (25 μL) is made up of 12.5 μL of 2× TaqMan universal PCR master mixture (PCR buffer, deoxynucleoside triphosphates, AmpliTaq Gold, an internal reference signal [6-carboxy-X-rhodamine], uracil N-glycosylase, MgCl2; Applied Biosystems), 300 nM forwar primer, 300 nM reverse primer, 300  nM probe and 5 μL of purified DNA from plaque samples. The negative control is 5 μL of sterile H2O. 2. The mixture is subjected to an initial amplification cycle of 50°C for 2 min and 95°C for 10 min, followed by 45 cycles at 95°C for 15 s and 60°C for 1 min. The data are analyzed with ABI 7000  Sequence Detection System software. 3. The analytical sensitivity of each of the five real-time PCR sets is determined in triplicate with DNA isolated from Fusobacterium strain. Serial tenfold dilutions of homologous DNA are used as standard curves. Note: The Ct parameter is defined as the fractional cycle number at which the fluorescence of the reporter dye generated by cleavage of the probe crosses an arbitrarily defined threshold within the logarithmic phase. This parameter can be used to compare different amplification reactions. (ii) Real-Time PCR for Fusobacterium spp.  Martin et al.73 reported a real-time PCR for Fusobacterium spp. with primers and probe from a conserved region in the 16S rRNA gene (rDNA) sequence (Table 48.5). The probe is labeled with fluorescent dyes 6-carboxyfluorescein (FAM) at the 5′ end and 6-carboxytetramethylrhodamine (TAMRA) at the 3′ end. Procedure 1. Bacteria are cultured to the late exponential phase, harvested by centrifugation (14,000 × g at 18 to 20°C for 2 min), washed, and resuspended in 10 mM phosphate buffer pH 6.7 containing 1 mg of lysozyme/mL, 1 mg of mutanolysin/mL, and 5 mM ZnCl2. After incubation at 60°C for 30 min, DNA is

TABLE 48.4 Primers and Fluorogenic Probes for the Specific Detection of Fusobacterium spp. Primer or Probe Forward Reverse Probe

Sequence (5′→3′) GGATTTATTGGGCGTAAAGC GGCATTCCTACAAATATCTACGAA FAM-CTCTACACTTGTAGTTCCG-TAMRA

Product (bp) 162

550

Molecular Detection of Human Bacterial Pathogens

TABLE 48.5 Sequences of Oligonucleotide Primers and Probe for Fusobacterium

TABLE 48.6 Primers and Probe for the Real-Time PCR Detection of Fusobacterium spp.

Primer or Probea

Primer or Probe

Forward Reverse Probe

a









Sequence (5′ → 3′)

Tm (°C)

AAGCGCGTCTAGGTGGTTATGT TGTAGTTCCGCTTACCTCTCCAG FAM-CAACGCAATACAGAGTTGAGCCCTG CATT-TAMRA

58.8 58.6 69.9

Fs619F CGCAGAAGGTGAAAGTCCTGTAT Fs719R TGGTCCTCACTGATTCACACAGA Probe Fs663T FAM-ACTTTGCTCCCAAGTAACATGG AACACGAG-TAMRA

The Fusobacterium-specific primer and probe set also detects F. periodonticum, F. alocis, and F. simiae if present.

extracted and purified with a QIAamp DNA Mini Kit (QIAGEN). The DNA concentration (A260) and purity (A260/A280) are measured. 2. To extract DNA from the anaerobic bacteria present in homogenized carious dentine, frozen suspensions are thawed on ice, and 80-μL samples are combined with 100 μL of ATL buffer (QIAGEN) and 400 μg of proteinase K (QIAGEN). The samples are vortexed for 10 s prior to incubation at 56°C for 40 min, with vortexing every 10 min to lyze the cells. Following the addition of 200 μg of RNase (Sigma), the samples are incubated for a further 10 min at 37°C before the DNA is purified with a QIAamp DNA Mini Kit. 3. Real-time PCR mixture (25-μL) is prepared with the TaqMan Universal PCR Master Mix containing 100 nM each forward primer, reverse primer, and probe and between 10 and 100 pg of template DNA/μL. 4. The real-time PCR amplification is conducted at 50°C for 2 min and 95°C for 10 min, followed by 40 cycles at 95°C for 15 s and 60°C for 1 min in an ABI PRISM 7700 Sequence Detection System (Applied Biosystems). Data analysis is performed with Sequence Detection software (version 1.6.3, Applied Biosystems). 5. The sensitivity of real-time PCR in detecting DNA is determined in duplicate with DNA extracted from Fusobacterium strain by using the appropriate homologous DNA as a standard.

(iii) Real-Time PCR for Fusobacterium spp.  Suzuki et al.55 described a quantitative real-time PCR targeting the 16S rRNA gene for detection of Fusobacterium spp. in subgingival plaque samples (Table 48.6). Procedure 1. Real-time PCR mixture (20-μL) is composed of 1 μL of lyzed cells, 1× qPCR Master Mix (Eurogentec), sense and antisense primer at 200 nM each, and the TaqMan probe at 250 nM. 2. Amplification is conducted at 50°C for 2 min and 95°C for 10 min, followed by 60 cycles of 95°C for

Sequence (5′→3′)



Product (bp) 101

15 s and 58°C for 1 min with the ABI PRISM 7700 Sequence Detection System (Applied Biosystems). 3. Standard curve is plotted with the Ct values obtained by amplifying successive tenfold dilutions of a known concentration of DNA extracted from bacterial cells containing 1.2 × 106 CFU (230 ng/ μL) for F. nucleatum ATCC 10953.

(iv) Sequencing Analysis of Fusobacterium 16S rRNA Gene  Cahill et al.69 utilized the conserved DNA sequences flanking the regions of DNA variation in the 16S rRNA gene to design degenerate primers 16S2F (5′-CCTACGGRSGCAGCAG-3′, positions 299–320) and 16S2R (5′-GGACTACCMGGGNTATCTAATCCKG-3′, positions 741–764) for identification of Fusobacterium spp. from fetal membranes (amnion, chorion, and adherent decidua). Procedure 1. Fetal membranes (amnion, chorion, and adherent decidua) are obtained within 30 min of delivery and placed in phosphate-buffered saline (PBS) supplemented with penicillin-streptomycin to limit further bacterial growth. Tissue samples (~200 mg) are frozen in liquid nitrogen and kept at −80°C until further study. DNA is extracted from the membrane tissue using the DNeasy Tissue Extraction Kit (Qiagen), resuspended in 200 µL of double distilled H2O and stored at −20°C. 2. PCR mixture (50 μL) is composed of 1.5 mM MgCl2, 50 mM KCl, 10 mM Tris-HCl pH 8.3, 200 µM of each deoxynucleotide triphosphate, 2 U of Amplitaq DNA polymerase (PE Applied Biosystems) and extracted DNA. 3. Amplification is performed in an ABgene® thermocycler with 95°C for 1 min, 35 cycles of 95°C for 30 s, 50°C for 30 s, 72°C for 1 min, and a final 72°C for 5 min. Negative control is included in each run with double distilled H2O replacing DNA. 4. PCR products are resolved by agarose gel electrophoresis and the amplicon of required size is extracted using the Qiaex II gel extraction kit (Qiagen). The resulting DNA is ligated into the PCR®-TOPO® TA vector (Invitrogen) and transformed into One Shot® Top 10 chemically competent E. coli (Invitrogen).

Fusobacterium

Twelve clones per sample are sequenced. All sequencing results obtained are analyzed using the BLAST program and the resulting alignments analyzed. Note: The integrity of the human DNA from these extractions may be confirmed by amplification of β -actin using specific primers Actin F and Actin R.69 F. nucleatum is found to be the most commonly detected organism, being present in 9 of the 15 samples (60%) that contained bacterial DNA.69

551

48.3  CONCLUSION Members of the genus Fusobacterium are gram-positive, nonspore-forming, nonmotile, anaerobic, rods that are present on the mucosal surfaces (e.g., mouth, upper respiratory tract, gastrointestinal tract, female genital tract) of humans and animals as commensal microflora.21,74 Several Fusobacterium species/subspecies such as F. necrophorum and F. nucleatum have the capacity to cause various purulent or gangrenous infections in humans, particularly after accidental trauma, surgery, and edema.75 While phenotypic identification of Fusobacterium spp. using cell and colony morphology as well as biochemical properties is possible, it is far too time consuming and complex to be of value for guiding the treatment and management of the infection.21,76–78 The development of nucleic acid–amplification technologies such as PCR has helped streamline the identification and detection of Fusobacterium species/subspecies. Use of PCR assays has not only facilitated rapid, sensitive, and specific differentiation of Fusobacterium bacteria causing diseases in humans and animals, but also revealed insights in the molecular evolution and epidemiology of these organisms, leading to implementation of improved control and prevention measures against infections due to Fusobacterium spp.









REFERENCES



1. Dorsch, M., Love, D.N., and Bailey, G.D. Fusobacterium equinum sp. nov., from the oral cavity of horses, Int. J. Syst. Evol. Microbiol., 51, 1959, 2001. 2. Batty, A., Wren, M.W., and Gal, M., Fusobacterium necrophorum as the cause of recurrent sore throat: Comparison of isolates from persistent sore throat syndrome and Lemierre’s disease, J. Infect., 51, 299, 2004. 3. Nagaraja, T.G. et al., Fusobacterium necrophorum infections in animals: Pathogenesis and pathogenic mechanisms, Anaerobe, 11, 239, 2005. 4. Bennett, G. et al., Detection of Fusobacterium necrophorum and Dichelobacter nodosus in lame cattle on dairy farms in New Zealand, Res. Vet. Sci., 87, 413, 2009. 5. Altshuler, G., and Hyde, S., Clinicopathologic considerations of fusobacteria chorioamnionitis, Acta Obstet. Gynecol. Scand., 67, 513, 1988. 6. Albandar, J.M., Brown, L.J., and Loe, H., Putative periodontal pathogens in subgingival plaque of young adults with and without early-onset periodontitis, J. Periodontol., 68, 973, 1997.





7. Bourgault, A.-M., et al., Fusobacterium bacteremia: Clinical experience with 40 cases, Clin. Infect. Dis., 25, S181, 1997. 8. Brazier, J.S., Fusobacterium necrophorum infections in man, Rev. Med. Microbiol., 13, 141, 2002. 9. Berger, A. et al., Microbial invasion of the amniotic cavity at birth is associated with adverse short-term outcome of preterm infants, J. Perinat. Med., 31, 115, 2003. 10. Carta, G. et al., Periodontal disease and poor obstetrical outcome, Clin. Exp. Obstet. Gynecol., 31, 47, 2004. 11. Gardella, C. et al., Identification and sequencing of bacterial rDNAs in culture-negative amniotic fluid from women in premature labor, Am. J. Perinatol., 21, 319, 2004. 12. Goepfert, A.R. et al., Periodontal disease and upper genital tract inflammation in early spontaneous preterm birth, Obstet. Gynecol., 104, 777, 2004. 13. Shinjo, T., Fujisawa, T., and Mitsuoka, T., Proposal of two subspecies of Fusobacterium necrophorum (Flügge) Moore and Holdeman: Fusobacterium necrophorum subsp. necrophorum subsp. nov., nom. rev. (ex Flügge 1886), and Fusobacterium necrophorum subsp. funduliforme subsp. nov., nom. rev. (ex Halle 1898), Int. J. Syst. Bacteriol., 41, 395, 1991. 14. Dzink, J.L., Sheenan, M.T., and Socransky, S.S., Proposal of three subspecies of Fusobacterium nucleatum Knorr 1922: Fusobacterium nucleatum subsp. nucleatum subsp. nov., comb. nov.; Fusobacterium nucleatum subsp. polymorphum subsp. nov., nom. rev., comb. nov.; and Fusobacterium nucleatum subsp. vincentii subsp. nov., nom. rev., comb. nov., Int. J. Syst. Bacteriol., 40, 74, 1990. 15. Gharbia, S.E., and Shah, H.N., Fusobacterium nucleatum subsp. fusiforme subsp. nov. and Fusobacterium nucleatum subsp. animalis subsp. nov. as additional subspecies within Fusobacterium nucleatum, Int. J. Syst. Bacteriol., 42, 296, 1992. 16. Kapatral, V. et al., Genome sequence and analysis of the oral bacterium Fusobacterium nucleatum strain ATCC 25586, J. Bacteriol., 184, 2005, 2002. 17. Karpathy, S.E. et al., Genome sequence of Fusobacterium nucleatum subspecies polymorphum—a genetically tractable fusobacterium, PLoS One, 2, e659, 2007. 18. Periasamy, S., and Kolenbrander, P.E., Aggregatibacter actinomycetemcomitans builds mutualistic biofilm communities with Fusobacterium nucleatum and Veillonella species in saliva, Infect. Immun., 77, 3542, 2009. 19. Roberts, G.L., Fusobacterial infections: An underestimated threat, Br. J. Biomed. Sci., 57, 156, 2000. 20. Ludlam, H. et al., Epidemiology of pharyngeal carriage of Fusobacterium necrophorum, J. Med. Microbiol., 58, 1264, 2009. 21. Riordan, T., Human infection with Fusobacterium necrophorum (necrobacillosis), with a focus on Lemierre’s syndrome, Clin. Microbiol. Rev., 20, 622, 2007. 22. Hagelskjaer, L.H. et al., Incidence and clinical epidemiology of necrobacillosis, including Lemierre’s syndrome, in Denmark 1990–1995, Eur. J. Clin. Microbiol. Infect. Dis., 17, 561, 1998. 23. Hagelskjaer Kristensen, L., and Prag, J., Human necrobacillosis, with emphasis on Lemierre’s syndrome, Clin. Infect. Dis., 31, 524, 2000. 24. Karkos, P.D. et al., Lemierre’s syndrome: A systematic review, Laryngoscope, 119, 1552, 2009. 25. Narayanan, S.K. et al., Cloning, sequencing, and expression of the leukotoxin gene from Fusobacterium necrophorum, Infect. Immun., 69, 5447, 2001.

552 26. Narayanan, S.K. et al., Leukotoxins of gram-negative bacteria, Vet. Microbiol., 84, 337, 2002. 27. Narayanan, S.K. et al., Fusobacterium necrophorum leukotoxin induces activation and apoptosis of bovine leukocytes, Infect. Immun., 70, 4609, 2002. 28. Tan, Z.L., Nagaraja, T.G., and Chengappa, M.M., Factors affecting the leukotoxin activity of Fusobacterium necrophorum, Vet. Microbiol., 32, 15, 1992. 29. Tan, Z.L., Nagaraja, T.G., and Chengappa, M.M., Fusobacterium necrophorum infections: Virulence factors, pathogenic mechanism and control measures, Vet. Res. Commun., 20, 113, 1996. 30. Ludlam, H.A. et al., lktA-encoded leukotoxin is not a universal virulence factor in invasive Fusobacterium necrophorum infections in animals and man, J. Med. Microbiol., 58, 529, 2009. 31. Oelke, A.M. et al., The leukotoxin operon of Fusobacterium necrophorum is not present in other species of Fusobacterium, Anaerobe, 11, 123, 2005. 32. Zhang, F. et al., The two major subspecies of Fusobacterium necrophorum have distinct leukotoxin operon promoter regions, Vet. Microbiol., 112, 73, 2006. 33. Tadepalli, S. et al., Human Fusobacterium necrophorum strains have a leukotoxin gene and exhibit leukotoxic activity, J. Med. Microbiol., 57, 225, 2008. 34. Feuille, F.L. et al., Synergistic tissue destruction induced by Porphyromonas gingivalis and Fusobacterium nucleatum, J. Dent. Res., 73, 159, 1994. 35. Bradshaw, D.J. et al., Role of Fusobacterium nucleatum and coaggregation in anaerobe survival in planktonic and biofilm oral microbial communities during aeration, Infect. Immun., 66, 4729, 1998. 36. Han, Y.W. et al., Interactions between periodontal bacteria and human oral epithelial cells: Fusobacterium nucleatum adheres to and invades epithelial cells, Infect. Immun., 68, 3140, 2000. 37. Han, Y.W. et al., Fusobacterium nucleatum induces premature and term stillbirths in pregnant mice: Implication of oral bacteria in preterm birth, Infect. Immun., 72, 2272, 2004. 38. Kulekci, G. et al., PCR analysis of Actinobacillus actinomycetemcomitans, Porphyromonas gingivalis, Treponema denticola and Fusobacterium nucleatum in middle ear effusion, Anaerobe, 7, 241, 2001. 39. Moraes, S.R. et al., Clonality of Fusobacterium nucleatum in root canal infections, Oral Microbiol. Immunol., 17, 394, 2002. 40. Nyfors, S. et al., Emergence of penicillin resistance among Fusobacterium nucleatum populations of commensal oral flora during early childhood, J. Antimicrob. Chemother., 51, 107, 2003. 41. Haraldsson, G., Holbrook, W.P., and Könönen, E., Clonal similarity of salivary and nasopharyngeal Fusobacterium nucleatum in infants with acute otitis media experience, J. Med. Microbiol., 53, 161, 2004. 42. Haraldsson, G., Holbrook, W.P., and Könönen, E., Clonal persistence of oral Fusobacterium nucleatum in infancy, J. Dent. Res., 83, 500, 2004. 43. Thurnheer, T. et al., Infinite serovar and ribotype heterogeneity among oral Fusobacterium nucleatum strains? Anaerobe, 5, 79, 1999. 44. Diaz, P.I., Zilm, P.S., and Rogers, A.H., Fusobacterium nucleatum supports the growth of Porphyromonas gingivalis in oxygenated and carbon-dioxide-depleted environments, Microbiology, 148, 467, 2002.

Molecular Detection of Human Bacterial Pathogens 45. Tew, J.G., Thomas, S.S., and Ranney, R.R., Fusobacterium nucleatum-mediated immunomodulation of the in vitro secondary antibody response to tetanus toxoid and Actinobacillus actinomycetemcomitans, J. Periodont. Res., 22, 506, 1987. 46. Gaetti-Jardim Junior, E., and Avila-Campos, M.J., Haemagglutination and haemolysis by oral Fusobacterium nucleatum, New Microbiol., 22, 63, 1999. 47. Jensen, A., Hagelskjaer Kristensen, L., and Prag, J., Detection of Fusobacterium necrophorum subsp. funduliforme in tonsillitis in young adults by real-time PCR, Clin. Microbiol. Infect., 13, 695, 2007. 48. Hall, V. et al., A comparative study of Fusobacterium necrophorum strains from human and animal sources by phenotypic reactions, pyrolysis mass spectrometry and SDS-PAGE, J. Med. Microbiol., 46, 865, 1997. 49. Brown, R., Lough, H.G., and Poxton, I.R., Phenotypic characteristics and lipopolysaccharides of human and animal isolates of Fusobacterium necrophorum, J. Med. Microbiol., 46, 873, 1997. 50. Ashimoto, A. et al., Polymerase chain reaction detection of 8 putative periodontal pathogens in subgingival plaque of gingivitis and advanced periodontitis lesions, Oral Microbiol. Immunol., 11, 266, 1996. 51. George, K.S., Reynolds, M.A., and Falkler, W.A. Jr., Arbitrarily primed polymerase chain reaction fingerprinting and clonal analysis of oral Fusobacterium nucleatum isolates, Oral Microbiol. Immunol., 12, 219, 1997. 52. Suchett-Kaye, G., Décoret, D., and Barsotti, O., Clonal analysis by ribotyping of Fusobacterium nucleatum isolates obtained from healthy young adults with optimal plaque control, J. Periodont. Res., 33, 179, 1998. 53. Jin, J. et al., Characterization of the 16S−23S rRNA intergenic spacer regions among strains of the Fusobacterium necrophorum cluster, J. Vet. Med. Sci., 64, 273, 2002. 54. Narongwanichgarn, W. et al., Specific detection and differentiation of two subspecies of Fusobacterium necrophorum by PCR, Vet. Microbiol., 91, 183, 2003. 55. Suzuki, N. et al., Quantitative microbiological study of subgingival plaque by real-time PCR shows correlation between levels of Tannerella forsythensis and Fusobacterium spp., J. Clin. Microbiol., 42, 2255, 2004. 56. Jervøe-Storm, P.M. et al., Comparison of culture and real-time PCR for detection and quantification of five putative periodontopathogenic bacteria in subgingival plaque samples, J. Clin. Periodontol., 32, 778, 2005. 57. Kim, H.S. et al., Development of strain-specific PCR primers based on a DNA probe Fu12 for the identification of Fusobacterium nucleatum subsp. nucleatum ATCC 25586T, J. Microbiol., 43, 331, 2005. 58. Cogulu, D. et al., PCR-based identification of selected pathogens associated with endodontic infections in deciduous and permanent teeth, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 106, 443, 2008. 59. Rôças, I.N., and Siqueira, J.F. Jr., Identification of bacteria enduring endodontic treatment procedures by a combined reverse transcriptase–polymerase chain reaction and reversecapture checkerboard approach, J. Endodontics, 36, 45, 2010. 60. Greisen, K. et al., PCR primers and probes for the 16S rRNA gene of most species of pathogenic bacteria, including bacteria found in cerebrospinal fluid, J. Clin. Microbiol., 32, 335, 1994. 61. Jalava, J. et al., Bacterial 16S rDNA polymerase chain reaction in the detection of intra-amniotic infection, Br. J. Obstet. Gynaecol., 103,664, 1996.

Fusobacterium 62. Hitti, J. et al., Broad-spectrum bacterial rDNA polymerase chain reaction assay for detecting amniotic fluid infection among women in premature labor, Clin. Infect. Dis., 24, 1228, 1997. 63. Jin, J. et al., Comparison of the 16S-23S rRNA intergenic spacer regions between Fusobacterium varium and “Fusobacterium pseudonecrophorum” strains, J. Vet. Med. Sci., 64, 285, 2002. 64. Aliyu, S.H. et al., Real-time PCR investigation into the importance of Fusobacterium necrophorum as a cause of acute pharyngitis in general practice, J. Med. Microbiol., 53, 1029, 2004. 65. Jin, J. et al., Phylogenetic analysis of Fusobacterium necrophorum, Fusobacterium varium and Fusobacterium nucleatum based on gyrB gene sequences, J. Vet. Med. Sci., 66, 1243, 2004. 66. Avila-Campos, M.J. et al., Arbitrarily primed-polymerase chain reaction for identification and epidemiologic subtyping of oral isolates of Fusobacterium nucleatum, J. Periodontol., 70, 1202, 1999. 67. Narongwanichgarn, W. et al., Differentiation of Fusobacterium necrophorum subspecies from bovine pathological lesions by RAPD-PCR, Vet. Microbiol., 82, 383, 2001. 68. Rinttila, T. et al., Development of an extensive set of 16S rDNA-targeted primers for quantification of pathogenic and indigenous bacteria in faecal samples by real-time PCR, J. Appl. Microbiol., 97, 1166, 2004. 69. Cahill, R.J. et al., Universal DNA primers amplify bacterial DNA from human fetal membranes and link Fusobacterium nucleatum with prolonged preterm membrane rupture, Mol. Hum. Reprod., 11, 761, 2005.

553 70. D’Ercole, S. et al., Comparison of culture methods and multiplex PCR for the detection of periodontopathogenic bacteria in biofilm associated with severe forms of periodontitis, New Microbiol., 31, 383, 2008. 71. Shin, H.S. et al., Development of strain-specific PCR primers for the identification of Fusobacterium nucleatum subsp. fusiforme ATCC 51190(T) and subsp. vincentii ATCC 49256(T), Anaerobe, 16, 43, 2010. 72. Boutaga, K. et al., Periodontal pathogens: A quantitative comparison of anaerobic culture and real-time PCR, FEMS Immunol. Med. Microbiol., 45, 191, 2005. 73. Martin, F.E. et al., Quantitative microbiological study of human carious dentine by culture and real-time PCR: Association of anaerobes with histopathological changes in chronic pulpitis, J. Clin. Microbiol., 40, 1698, 2002. 74. Batty, A., and Wren, M.W., Prevalence of Fusobacterium necrophorum and other upper respiratory tract pathogens isolated from throat swabs, Br. J. Biomed. Sci., 62, 66, 2005. 75. Smith, G.R., and Thornton, E.A., Pathogenicity of Fusobacterium necrophorum strains from man and animals, Epidemiol. Infect., 110, 499, 1993. 76. Hill, G.B., Preterm birth: Associations with genital and possibly oral microflora, Ann. Periodontol., 3, 222, 1998. 77. Tan, Z.L. et al., Biological and biochemical characterization of Fusobacterium necrophorum leukotoxin, Am. J. Vet. Res., 55, 515, 1994. 78. Narayanan, S. et al., Ribotyping to compare Fusobacterium necrophorum isolates from bovine liver abscesses, ruminal walls, and ruminal contents, Appl. Environ. Microbiol., 63, 4671, 1997.

and Leptotrichia49 Leptotrichia Like Organisms Emenike Ribs K. Eribe and Ingar Olsen CONTENTS 49.1 Introduction...................................................................................................................................................................... 555 49.1.1 Classification and Morphology............................................................................................................................. 555 49.1.2 Biology and Epidemiology................................................................................................................................... 556 49.2.3 Clinical Features and Pathogenesis...................................................................................................................... 558 49.1.4 Diagnosis.............................................................................................................................................................. 559 49.2 Methods............................................................................................................................................................................ 560 49.2.1 Sample Preparation............................................................................................................................................... 560 49.2.2 Detection Procedures............................................................................................................................................ 562 49.3 Conclusion and Future Perspectives................................................................................................................................. 563 Acknowledgments...................................................................................................................................................................... 563 References.................................................................................................................................................................................. 563

49.1  INTRODUCTION 49.1.1  Classification and Morphology Classification. Trevisan1 established the genus Leptotrichia in 1879 to distinguish filamentous microorganisms in the oral cavity from free-living filamentous Leptothrix species. The major metabolic end product of Leptotrichia,2 lactic acid, separates it from other filamentous bacteria, Fusobacterium and Bacteroides.3 In the draft taxonomic outline4 based on comparative analysis of 16S rRNA sequences, Leptotrichia is proposed as the first genus in the family Leptotrichiaceae, phylum Fusobacteria. Other proposed members of this family are the genera Sebaldella, Streptobacillus, and Sneathia.4 Molecular microbiological studies have revealed a wide biodiversity in Leptotrichia.5,6 Most of these organisms cannot be cultured. A large variety of new 16S rRNA phylotypes has been detected in Leptotrichia from humans5,7,8 and animals9,10 as well as from environmental samples (http://www.meg. boun.edu.tr/MKC.pdf). The genus Leptotrichia contains six validated species: L. buccalis, which is the type species; L. goodfellowii, L. hofstadii, L. shahii, L. trevisanii,3,11,12 and L. wadei.5,11 “L. amnionii”13 has not been formally validated but should probably be transferred to the genus Sneathia.5 The G + C content of DNA in Leptotrichia is 25 mol%14 or 24–29.7 mol%.12,15 DNA–DNA hybridization showed more than 70% homology between all species of Leptotrichia. With L. buccalis, L. goodfellowii showed 3.8%–5.5% DNA–DNA relatedness; L. shahii showed 24.5%–34.1% relatedness;

L. hofstadii exhibited 27.3%–36.3% relatedness; and L. wadei 24.1%–35.9% relatedness, respectively.5 Leptotrichia is generally considered negative with respect to catalase, oxidase, and indole production. Some species are catalase-positive5 (Table 49.1). L. buccalis hydrolyses aesculin.2,5,16 Classification of Leptotrichia involves not only genetic and biochemical analyses but also fatty acid and protein profiles (see later). Morphology. Leptotrichia are straight or curved  rods, 0.8–1.5 µm wide and 5–15 µm long, nonsporing and non­ motile5,15 with one or both ends pointed or rounded. Some features of Leptotrichia species are also shown in Table 49.1. Cellular morphology of the novel species L. goodfellowii, L. hofstadii, L. shahii and L. wadei was studied by examining cells grown anaerobically at 37°C on Columbia or brainheart infusion (BHI) blood agar plates.5 Light microscopy (LM), transmission electron microscopy (TEM) and scanning electron microscopy (SEM) were used for characterization of cell morphology and ultrastructure. Leptotrichia species had several common features. Cells are arranged in pairs, some slightly curved, others in chains joined by flattened ends. Electron micrographs of ultrathin sections revealed elongated cells with a cell-wall structure typical of gram-negative bacteria. Leptotrichia possess a double plasma membrane layer, a single electron-dense (intermediate) layer and a double outer layer with scale-like protrusions on the surface referred to as “fish scales” (Figure 49.1).5,17–19 Fimbriae are not present. Septum is formed from the cytoplasmic membrane and the intermediate layer.5,18 The outer 555

556

Molecular Detection of Human Bacterial Pathogens

TABLE 49.1 Characteristics of Leptotrichia and Leptotrichia-Like Bacteria Character

L. buccalis

L. goodfellowii

L. hofstadii

L. shahii

L. wadei

L. trevisanii

Gram-negative Gram-negative Gram-negative

Colony morphology on blood agar

Smooth, rhizoid, or convoluted

Speckled, convex, irregular pink periphery, grayish light brown glistening surface, opaque, dry

Smooth, grayish, Smooth, low convex, convex, erose edged, “flecked” dark central spot with transillumi­ nation, slightly convoluted

Flat, rough, spreading margin,c adherent or low-convex, smooth, pigmented

Growth in air and 5%–10% CO2 on blood agar Indole Catalase Aesculin

+

+

Glistening, Very Convex, sparsely granular filamentous to filamentous to surface, rhizoid or irregular, old opaque, dry convoluted, colonies and circular, pale speckled, grayish brown convex, entire grayish with a with a dark (some dark central central spot, irregular and spot in old glistening, lobate), colonies, smooth with a grayish with a opaque, rough edge, dark central semidry in opaque, dry in spot consistency consistency + + +

+

−

+c

+ − +

− + +

− + +

− + +

− + +

− + UKd

+ + −

− − +

β-hemolytic

β-hemolytic

Nonhemolytic

β-hemolytic

UK

UK

UK

Lactic

Lactic

Lactic

Lactic

Lactic

Butyric

Acetic, succinic

Hemolysis on Nonhemolytic human blood agar Main fatty acids Lactic from glucosepeptone a b c d

7.5–15 µm long 7.5–17.5 µm rods long rods

Gram-negative

F. nucleatum Capnocytophaga

Cellular Gram-negative Gram-negative morphology Cells on bloodb 5–15 µm long, 2–4 µm short agar thick fusiform rods rods a

2.5–10 µm short 6–13 µm long rods fusiform rod

Gram-negative Gram-negative 3–10 µm long, 3–6 µm long, slender slender fusiform rods fusiform rods

Occasionally gram-positive in young cultures. Human blood. Gliding motility. Unknown.

membrane ridge is firmly attached to the peptidoglycan sacculus, which may be the point of origin of the structure. When analyzed by SDS-PAGE, the macromolecules forming the ridge revealed a major component of 210-kDa polypeptide and a 15-kDa minor component. It has been suggested that the 210-kDa molecule is likely an adhesion component, but this is not proven.18 A cross-section of the cell showed storage granules. Leptotrichia are usually anaerobic on first isolation with some strains growing aerobically in presence of CO2. The colonies on blood agar plates tend to be smooth, rhizoid, or convoluted and can also be sparsely filamentous to irregular and grayish brown in color, with a dark central spot in old colonies.3 Occasionally, colonies are opaque and dry in consistency.5

49.1.2  Biology and Epidemiology Leptotrichia are part of the normal flora in the oral cavity,20 intestine,15,21–29 and human female genitalia.22,26,30 Prior to tooth eruption, 17% of children harbor Leptotrichia, and it continues to be present in up to 71% after eruption.20,31 Leptotrichia can infect humans and animals. It has been reported to participate in oral diseases and diseases elsewhere in the body,3 for example, in the intestine15,21–29,32 and female genitalia22,26,30 of healthy persons. Leptotrichia has also been recovered from patients with gingivitis, necrotizing ulcerative gingivitis, adult/juvenile periodontitis, “refractory” periodontitis,33 noma,3,7,22,34–38 aortic aneurysms,8 cellulitis,39 phagedenic chancroid,40 salpingitis,41 acute appendicitis,42 and bacterial vaginosis.43 Children with gingivitis harbor three

Leptotrichia and Leptotrichia-Like Organisms

557

SLP

(a)

IDL

OM

(b) IM

G

G

IDL OM IM

(c) OM IM

SLP

(d)

G

IDL

(II)

G

S

SLP

SLP

(I)

FIGURE 49.1  TEM (a, b, c, and d) of cells of novel Leptotrichia species after anaerobic culture on Columbia blood agar at 37°C for 48 h. (a), (b), (c), and (d) Cross-sectional views of cells of L. goodfellowii LB 57T (a), L. hofstadii LB 23T (b), L. shahii LB 37T (c), and L. wadei LB 16T (d). In (d) cells are visible in both a longitudinal (I) and a cross-sectional (II) view. G, Granule; IDL, intermediate dense layer; IM, inner cytoplasmic membrane; OM, outer membrane; S, septum; SLP, scale-like protrusion; CM, cytoplasmic membrane. Bars, 0.1 (a, c) and 0.25 (b, d) µm. (Modified from Eribe et al., Int. J. Syst. Evol. Microbiol., 54, 583, 2004. With permission from the Society for Microbiology.)

times as many Leptotrichia species in subgingival plaque as adults with gingivitis.44 Leptotrichia has also been reported to colonize permucosal implants of edentulous patients.45 L. buccalis is highly saccharolytic, producing lactic acid, a property that may implicate its participation in tooth decay.46–49 The organism has also been recovered from peritoneal fluid50 and from blood of immunocompromised patients with neutropenia, HIV, leukemia, and endocarditis.34,36,39,46,50–60 Furthermore, Leptotrichia has been isolated from infected material of domestic animals and after dog bites.9,10 Most Leptotrichia species have been isolated from blood and characterized as L. buccalis, L. trevisanii, L. wadei, and L.  goodfellowii, whereas organisms such as L. hofstadii, L.  buccalis, and L. wadei have been recovered from saliva. Human Leptotrichia infections have been reported to occur mostly in immunocompromised patients with predisposing factors, but there have also been incidences in healthy humans. There is no report of disease transmission between humans. Leptotrichia buccalis. L. buccalis is the type species of the genus Leptotrichia. The type strain is ATCC 14201T=NCTC 10249T. It is a common member of the oral microflora in humans and has been isolated from dental plaque and saliva.20,61 It is generally considered catalase-, oxidase- and indole-negative, and it produces lactate, but catalase production can occur. L. buccalis hydrolyses aesculin.2,5,16 The DNA base content is 25 mol% G + C.14 L. buccalis has previously been called Leptothrix buccalis,62 Fusobacterium plautivincentii or Fusobacterium fusiforme.21 Leptotrichia was first cultivated by Wherry and Oliver,63 who named it Leptotrichia

innominata. Thjøtta et al.64 and Böe and Thjötta65 gave the first adequate descriptions of L. buccalis based on the fact that it had many features in common with Fusobacterium and classified it as a gram-negative anaerobic bacterium. Hofstad and Selvig17 used TEM to study its cell wall and concluded that L. buccalis belongs to gram-negative bacteria. Leptotrichia goodfellowii. The type strain of L. goodfellowii is CCUG 32286T=CIP 107915T 5 and was originally isolated from the blood of an endocarditis patients.56 It is a gram-negative, nonspore-forming, nonmotile short rod, varying in length, with one end tapered. Cells are arranged in pairs, some slightly curved; others, in chains joined by flattened ends. Colonies grow best anaerobically and sparsely aerobically at 37°C but not at 25°C and 42°C. Colonies are speckled, convex, irregular, pink in the periphery, and grayish light brown in color. They have a glistening surface appearance, are opaque, dry, and show β-hemolysis. L. goodfellowii is aesculin- and catalase-positive. Oxidase and indole are not produced. Arginine dihydrolase, β-galactosidase, β-glucosidase, β-N-acetyl glucosaminidase, alkaline phosphatase, arginine arylamidase, leucine arylamidase, and histidine arylamidase are produced. Leptotrichia hofstadii. The type strain of L. hofstadii is CCUG 47504T=CIP 107917T,5 and it was originally isolated from the saliva of a healthy person. It is a gram-negative, nonspore-forming, nonmotile, long rod, varying in length, with one end tapered. Cells are arranged in pairs, some slightly curved; others, in chains joined by flattened ends. Colonies grow best anaerobically and sparsely aerobically at 37°C but

558

not at 25°C and 42°C. Colonies are circular, convex, entire (some are irregular and lobate), and grayish in color with a dark central spot in old colonies. They have a glistening and granular surface appearance, are opaque, dry, and nonhemolytic. L. hofstadii is aesculin- and catalase-positive. Oxidase and indole are not produced. β-Galactosidase-6-phosphate, α-glucosidase, β-glucosidase, and alkaline phosphatase are generated. Mannose is fermented. Leptotrichia shahii. L. shahii has as type strain CCUG 47503T=CIP 107916T 5 and was originally isolated from a patient with gingivitis. It is a gram-negative, nonsporeforming, nonmotile long rod, varying in length, with one end tapered. Cells are arranged in pairs, some slightly curved; others in chains joined by flattened ends. Colonies grow best anaerobically and sparsely aerobically at 25°C and 37°C but not at 42°C. Colonies are very filamentous to rhizoid or convoluted, pale speckled, and grayish in color with a dark central spot in old colonies. They are opaque, semidry in consistency and nonhemolytic. L. shahii is aesculin- and catalase-positive. Oxidase and indole are not produced. α-Glucosidase and α-arabinosidase are generated. Leptotrichia wadei. The type strain of L. wadei is CCUG 47505T=CIP 107918T 5 and was originally isolated from the saliva of a healthy person. It is a gram-negative, nonsporeforming, nonmotile, short rod, varying in length, with one end tapered. Cells, some slightly curved, are arranged in pairs; others, in chains joined by flattened ends. L. wadei grows sparsely aerobically at 37°C but not at 25°C and 42°C. Colonies are convex, sparsely filamentous to irregular, and grayish brown in color, with a dark central spot in old colonies. The surface appearance is glistening and smooth with a rough edge. Colonies are opaque, dry in consistency, and β-hemolytic. The organism is aesculin- and catalase-positive. Oxidase and indole are not produced. However, α-glucosidase and β-glucosidase are generated. Leptotrichia trevisanii. L. trevisanii is a thin, filamentous, nonmotile, aerotolerant, gram-negative bacterium isolated from the blood of a man with acute myeloid leukemia.12 The type strain of L. trevisanii is ATCC 700907T 12=DSM 22070T. The deposition in a second culture collection in a different country (DSM) should now make L. trevisanii a validly published organism.3,11 The organism is catalase-positive but indole-, oxidase-, and urease‑negative. It is saccharolytic and produces lactate. The DNA base composition is 29.7 mol% G + C. The 16S rRNA gene sequences of L. trevisanii showed 96% identity with those of the type strain of L. buccalis,12 and 97% sequence identity with those of “Leptotrichia” species.53 “Leptotrichia amnionii.” “L. amnionii”13 has pleomorphic coccobacillary, long, nonmotile, fusiform cells. Some cells are joined end to end in a filamentous form. This bacterium was isolated from the amniotic fluid of a pregnant woman with intrauterine fetal demise. “L. amnionii” grows anaerobically on blood agar. The isolate was characterized only by its 16S rRNA gene sequences, which showed 96% relationship to Sneathia sanguinegens. “L. amnionii” is not yet validly published or recognized in IJSEM. Therefore, the

Molecular Detection of Human Bacterial Pathogens

taxonomic status of this species is uncertain, but it should probably be transferred to the genus Sneathia.5

49.1.3  Clinical Features and Pathogenesis Clinical Features. Leptotrichia has been isolated from a variety of clinical lesions in man. The oral lesions comprise gingival hyperplasia,43 gingivitis, necrotizing ulcerative gingivitis,66 adult/juvenile periodontitis, “refractory” periodontitis,33 noma,3,7 root caries,49 infected root canals,67–69 and malodor.70 It has also been found on permucosal implants of edentulous patients.42 Besides the mouth, other major sites of primary Leptotrichia colonization include the genitourinary tract, where these organisms have been found in bacterial vaginosis,71–75 abortion,76,77 genitourinary tract infection,78 salpingitis,41 preterm labor,79 preterm birth,80 and amniotic fluid infection.13 Leptotrichia can also spread from primary sites of colonization, causing acute appendicitis,42 cellulitis,39 arthritis,81 phagedenic chancroid,40 and Lemierre syndrome.82 It can further disseminate to the bloodstream of bone marrow transplants52 and immunocompromised patients with and without neutropenia.36,39,46,52,53,55–59,82–84 Leptotrichia has also been isolated from delivery and postpartum bacteremia.84 Further, the organisms have been recovered from peritonitis50 and from a hepatic abscess54 and blood cultures from immunocompetent patients.54,85 It has also been isolated from intestinal obstruction,50 aortic valve disease,34,40,58 aortic aneurysms,8 endocarditis,8,34,51,56,59,83 and esoinophilic pustular folliculitis, called “Ofuji’s disease” (www.emedicine. com/derm/byname/Eosinophilic-Pustular-Folliculitis.htm). Furthermore, Leptotrichia has been detected in dog-bite lesions.9,10 Pathogenesis. Leptotrichia has protruding structures on the cell surface18 (Figure 49.1). These ridges are probably adhesion structures.19 Buccal epithelial cells, red blood cells, HeLa, and embryonic kidney cells bound to the hemagglutination fragments of L. buccalis, while saliva displayed hemagglutination-inhibition activity in a manner suggesting a binding-site interaction.86 The receptor sites on human erythrocytes for L. buccalis may be sugar in nature.87 These authors suggested that a salivary aggregating component may possess the same specific group with which L. buccalis reacts on human erythrocytes, and that the enhanced adherence of L. buccalis cells to saliva-coated enamel powder is due to a greater affinity of the organism for this component that adsorbs to human enamel powder. As a gram-negative bacterium Leptotrichia possesses a potent lipopolysaccharide (LPS) (endotoxin) that displays an O-antigen linked to lipid A. The latter may cause hemorrhage, fever, tumor necrosis, fatal shock, septicemia, and even abortion. These features have been associated with cases of Leptotrichia infections. Endotoxin obtained from L. buccalis had a potent biological activity88 in New Zealand white female rabbits. The parameters studied were LD50, febrile, and leukocytic responses, and the dermal Shwartzman reaction. The LD50 of L. buccalis was 93.15 μg/kg with 95% confidence limits of 66.44

Leptotrichia and Leptotrichia-Like Organisms

and 131.00 μg/kg. Formal probit analysis revealed that the L. buccalis endotoxin, compared to that of Escherichia coli, had a relative potency of 19.45%, with 95% confidence limits of 13.50% and 28.01%. L. buccalis endotoxin was pyrogenic when given intravenously in a quantity of 0.039 μg/ kg, whereas E. coli endotoxin was pyrogenic at 0.009 μg/kg. Parallel line bioassay methods applied to the febrile doseresponse relationship of the toxins showed that, compared to E. coli, L. buccalis had a potency of 11.47% with 95% confidence limits of 9.69% and 13.63%. The quantity of endotoxin necessary to elicit a fever index of 40 cm2 was estimated as 0.285 and 2.099 μg/kg for E. coli and L. buccalis endotoxins, respectively. A significant correlation was found between febrile response and fever index. LPS isolated by phenol-water extraction from L. buccalis were endotoxic by their ability to alter dermal reactivity to epinephrine, and serologically specific by hemagglutination and hemagglutination-inhibition tests.89 Furthermore, L. buccalis exerted a blastogenic response of lymphocytes in periodontitis patients, who were significantly more reactive than in dentulous and normal subjects.90 This suggested that L. buccalis and other gram-negative bacteria tested, and the host response they evoke, are associated with advanced periodontal destruction. Systemic release of endotoxin and proinflammatory mediators from infected host tissue may contribute directly or indirectly to the sepsis syndrome associated with Leptotrichia. L. buccalis is highly saccharolytic, producing lactic acid, a property that may implicate participation in tooth decay.47–49,91 There is evidence supporting that Leptotrichia species are emerging human pathogens.76 Opportunistic pathogens tend to cause disease when local or general predisposing factors are present. This also relates to Leptotrichia. Localized Leptotrichia infections are inclined to occur when immunological defense mechanisms are compromised, as has been seen in cases with neutropenia,39,46,52,53,55–59 cancer,55,92 diabetes,81,93 pneumonia,94 peritoneal dialysis,50 and Lemierre syndrome.82 Leptotrichia infections may be serious in hospitalized patients, particularly those with cancer, as observed with L. shahii and L. wadei,92 and with bone marrow transplantation.46,52,55 Leptotrichia is apt to cause disease in the immunocompetent host.54 Localized Leptotrichia infections can lead to generalized infections and, occasionally, to fatal bacteremia and endocardial arrest, as reported with L. goodfellowii.51,56 A recent study showed that L. buccalis can occur together with other fusiform bacteria, such as fusobacteria, Tannerella forsythia, and Capnocytophaga species, in gingivitis and acute necrotizing ulcerative gingivitis, with high prevalence.66 In other cases of multiple organism infection, Leptotrichia species were recovered together with Bacteroides,42 Mycobacterium bovis, Candida albicans, and cytomegalovirus and herpes virus.50 Superinfection with the anaerobes L. buccalis, Treponema vincentii, and Fusobacterium nucleatum, responsible for the massive ulceration, has been associated with phagedenic chancroid patients. These patients present large, rapidly spreading necrotic ulcerations, which

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may result in extensive destruction or formation of a urethral fistula.40 “L. amnionii” was associated with clinical characteristics that were consistent with bacterial vaginosis and bacterial vaginosis, defined by Nugent’s and Amsel’s criteria. This organism together with other fastidious bacteria was suspected of causing unrecognized infection, since none were associated with abnormal vaginal discharge.95 The protective function of the indigenous bacterial flora can be disrupted by broad-spectrum antibiotic therapy,9,12,23,44,53,55–57,82,96 resulting in Leptotrichia infection. Enhanced Leptotrichia proliferation and tissue invasion can culminate into bloodstream invasion and dissemination.36,54,85 This particularly occurs when the patient’s immune system is compromised.

49.1.4  Diagnosis Leptotrichia are gram-negative, non-sporing, anaerobic, saccharolytic rods. Fresh cells can occasionally stain grampositive. Usually, Leptotrichia are anaerobic on the first isolation, but some strains grow aerobically in presence of CO215 (Table 49.1). Some strains are aerotolerant. The colonies on blood agar plates are 0.5–3.0 mm in diameter. They tend to be smooth, rhizoid, or convoluted but can also be sparsely filamentous to irregular and grayish brown in color, with a dark central spot in old colonies.3 Occasionally, colonies are opaque and dry in consistency.5 For correct diagnosis of Leptotrichia, adequate facilities for anaerobic cultivation and qualified expertise in anaerobic microbiology and taxonomy are needed.5,6,97–100 L. buccalis is isolated from clinical specimens after anaerobic culture on blood agar such as Brucella blood agar or FFA supplemented with sheep or horse blood. To isolate L. buccalis from the normal flora, addition of josamycin, vancomycin, and norfloxacin to the blood agar medium has been recommended.15 Others have used Fusobacterium egg yolk agar for recovery of Leptotrichia.101 A selective medium for culture of fusobacteria and leptotrichia has been established.102 These authors found that Blood Agar Base (Difco), containing 5% defibrinated sheep blood in the natural or “chocolate” state with the addition of vancomycin (7.5 µg/mL of medium) and streptomycin (20 µg/mL), was an excellent medium for isolation of these bacteria. A reliable diagnosis of L. buccalis can be made from cellular and colony morphology and a few biochemical tests. Biochemical reactions, cellular fatty acid (CFA) composition, and lactic acid as the major end product of glucose metabolism were used to distinguish them from fastidious gramnegative rods thought to be Capnocytophaga species.103 Diagnosis of Leptotrichia should include both phenotypic and genetic methods.5,6,97 It is no doubt that Leptotrichia has been underdiagnosed in the past due to problems with distinguishing it from closely related organisms such as Fusobacterium, Capnocytophaga, and Lactobacillus, and the failure to cultivate many Leptotrichia strains.5 Useful in the presumptive identification of Leptotrichia may be their susceptibility to antibiotics. L. buccalis can

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be susceptible to β-lactamantibiotics, carbapenems, clindamycin, chloramphenicol, tetracycline, and metronidazole, but resistant to vancomycin and aminoglycosides.21,57 Most strains are moderately resistant or resistant to erythromycin. It is difficult to diagnose Leptotrichia infection based on clinical features alone. Fever can be an indication of LPS release associated with Leptotrichia infection. Also, clinical diagnosis of infection in the compromised host should cause consideration of Leptotrichia infection. It is important to realize that a number of Leptotrichia isolates have not yet been cultivated.6,104 Molecular techniques should therefore be used to reveal the complete spectrum of Leptotrichia infections in man.3

49.2  METHODS 49.2.1  Sample Preparation (i) Bacterial Culture and Phenotypic Characterization  The present methods, described by Eribe et al.5,6,97 have been used successfully for phenotypic and genetic characterization of Leptotrichia strains and for establishing new Leptotrichia species. For additional details and references, see these publications. Other methods may also be applicable, but the reported ones are working best in our hands. Cultivation of Bacterial Strains. Suspected Leptotrichia strains are seeded onto Trypticase soy agar plates supplemented with 5% human blood, 50 mg/mL of hemin, and 5 mg/mL of menadione. They are cultured anaerobically (90% N2, 5% H2, and 5% CO2) at 37°C for 2–5 days in evacuation jars (Anoxomat System, WS9000, Mart, The Netherlands). After replating, single colonies are picked and subcultured anaerobically on blood agar plates at 37°C for 2–3 days. After the culturing and harvesting of bacteria, cells are generally identified by gram staining, which is then followed by conventional phenotypic or genotypic procedures, or both. The isolated Leptotrichia strains may be identified by a commercial identification kit (rapid ID 32 A, bioMérieux), and CFA-, and protein-analyses. Alternatively or as a supplement, DNA is extracted from the isolates for RAPD, specific PCR, and 16S rRNA gene sequence analysis. A polyphasic approach secures good taxonomy. Enzymatic/Biochemical Testing. For enzymatic/biochemical testing the bacteria are cultured anaerobically at 37°C for 2–3 days on Columbia base blood agar supplemented with 5% human blood, 50 mg/mL of hemin, and 5 mg/mL of menadione. A bacterial suspension with a turbidity equivalent to a McFarland number 4 standard is made from the culture. Suspensions are transferred to the rapid ID 32 A kit (API), which is based on 29 standardized and miniaturized enzymatic and biochemical tests. The kits are filled using a robot inoculator and incubated anaerobically at 37°C for 4 h. Reading of the kits occurs automatically in an ATB reader. The results of the reactions are transferred into a numerical code and treated automatically in a database for Leptotrichia identification (API Plus). The experiments are done in triplicate, starting from the growth of the cultures through the analyses.

Molecular Detection of Human Bacterial Pathogens

Cellular Fatty Acid Extraction. For CFA analysis the Leptotrichia cultures are transferred from secondary blood agar plates to liquid glass tubes containing 10 mL of prereduced anaerobically sterilized (PRAS) peptone-yeastglucose (PYG) broth under anaerobic conditions, using a Virginia Polytechnic Institute and State University (VPI) (Blacksburg, VA) anaerobic culture system (Bellco, Inc., Vineland, NJ). The glass tubes are cultured anaerobically at 37°C for 2–5 days. After culturing, the bacterial cells are harvested by centrifugation, washed once in 0.85% NaCl and once in deionized distilled water, pelleted by centrifugation, and stored at −70°C until analyzed. Pelleted cells (1–5 mg) kept in 8-mL glass tubes with Teflon-lined screw caps, are thawed prior to extraction. Saponification, methylation, and extraction of the cellular fatty acids are performed as recommended in the operation manual of the Microbial Identification System (MIS) software package (Microbial Identification System, ID). The cells are lyzed and saponified with 1 M saponification reagent [sodium hydroxide, methanol (certificate grade), distilled water]. The tubes are closed and their contents mixed for 10 s using a whirly mixer, and then heated in a boiling water bath for 5 min. After remixing the contents, the tubes are heated in the water bath for additional 25 min. They are then cooled for 30–60 s in a tray with cold tap water. For methylation, HCl-methanol (6.0 N HCl, methanol and sulfuric acid–methanol [50% H2SO4 (certificate grade), methanol] are added and mixed for 10 s. The tubes are heated at 80°C for 10 min and then again cooled for 30–60 s in a tray with cold tap water. The methylated components are extracted by adding hexane-ether [hexane and methyl-tert-butyl ether (HPLC grade)]. During extraction the tubes are placed in a test tube rotator, and mixing of their contents occurs by rotating the tubes end-over-end for 10 min. After the contents have separated in two phases, the lower aqueous phase is collected and discarded, whereas the upper (organic) phase is retained. Each extract is washed with NaOH–NaCl (sodium hydroxide in deionized distilled water saturated with sodium chloride). The tubes are again rotated end-over-end for 5 min, and the contents allowed to settle down. By using a Pasteur pipette, two-thirds of the top phase are transferred to a 2.0-mL gas chromatographic sample vial [Hewlett-Packard (H-P)]. The vials are closed using 11 mm caps with a red silicone septum lined with Teflon (H-P) and labeled. The experiments are done in triplicate, including cultures, extraction, and gas chromatography. Cellular Fatty Acid Analysis by Gas Chromatography. CFA profiles are obtained using a model H-P-5980 series II gas chromatograph (H-P) equipped with an H-P U2 crosslinked 5% phenyl-methyl silicone fused silica capillary column (25 m × 0.2 m I.D.), a flame ionization detector, a model 3392A integrator, a model 7673A automatic sampler, and a computer 486DX with 16 MB of RAM. Hydrogen is used as carrier gas at 0.5 mL/min. The gas flow rates for the detector are approximately 40 mL/min for air, 30 mL/min for hydrogen, and 30 mL/min for nitrogen. The temperatures used are 250°C for the injection port and 300°C for the detector. After

Leptotrichia and Leptotrichia-Like Organisms

injection of 2 µL fatty acid extract, the oven temperature is increased from 170°C to 250°C at a rate of 5°C/min and then from 250°C to 300°C at a rate of 30°C /min, held at 300°C for 2 min and then returned to 170°C before the next sample is injected. Ninety-eight vials (49 bacterial strains in duplicate) are loaded onto a sample tray and analyzed. A calibration mixture (Calibration Standard Kit, Microbial ID) containing known fatty acids (straight-chain saturated C9:0-C20:0 fatty acid methyl esters and five hydroxy fatty acids (2-OHC10:0, 3-OH-C10:0, 2-OH-C14:0, 3-OH-C16:0, and 2-OH-C16:0), are included for quantitative assessment of the fatty acids, to compensate for peak area discrimination between low- and high-boiling-point fatty acids, and to provide accurate retention times for the straight-chain saturated fatty acids. The calibration mixture is analyzed before the fatty acid extracts and is automatically reanalyzed after every tenth injection. The quantitative data obtained from the fatty acid profiles are used as a basis for numerical analysis performed with assistance of the MIS software package (Microbial ID). The latter can identify the peaks (by retention time) and determine the area to height, the equivalent chain length (ECL), the total area, and the total area for named or listed compounds. The software package is used to calculate the percentage area for each named or listed compound compared to the total area of the compounds detected. These compounds are identified by using the Moore Broth Library, Version 3.9 (Microbial ID). Numerical Analysis of Fatty Acid Data. Numerical analysis of the quantitative fatty acid data obtained from the FAME profiles is made using the Library Generation Software, Version 1.0 (Microbial ID). Peak area values for each fatty acid are calculated as percentages of the total peak area to eliminate the effect of variation in inoculum size. Similarities are calculated and the coefficient based on the Euclidean distance between pairs of bacteria determined. Clustering of strains is achieved by the unweighted pair group method for arithmetic average with a program provided in the H-P Library Generation Software, resulting in a dendrogram. Culture for SDS-PAGE of Whole-Cell Proteins. For SDS-PAGE analysis of whole-cell protein, cultures are transferred from secondary plates to glass tubes containing 10 mL PRAS BHI broth (Biokar Diagnostics, Beauvais, France) supplemented with 0.1% calcium carbonate under anaerobic conditions (90% N2, 5% H2, 5% CO2). The organisms are cultured anaerobically at 37°C for 2–5 days in evacuation jars. After culturing, the bacterial cells are harvested by centrifugation at 13,000 × g for 4 min, washed once with 0.85% NaCl and deionized distilled water, and stored at −20°C or −86°C until analyzed. Whole-Cell Protein Extraction. Prior to the extraction of whole-cell proteins, the bacterial cells are thawed at room temperature and resuspended in TE buffer or in deionized distilled water. The cells are dispersed by sonication (Branson Ultrasonic Corporation) for 1 min at low amplification in an ice cooling bath. Acid-washed 106 µm glass beads are added to the tubes. The bacteria are vortexed twice, each time for 1 min, while kept on ice. From the vortexed bacterial

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suspension, 20 µL are transferred to Eppendorf tubes and 20  µL of treatment buffer consisting of stacking gel buffer (0.5 M Tris-HCl, pH 6.8, 10% SDS (w/v); 10% glycerol (w/v); 5% 2-mercaptoethanol (w/v); and 5 µL of loading dye (5 mg dissolved and filtered phenol red) are added. The mixture is boiled for 5 min, centrifuged at 13, 000 × g for 5 min and then kept on ice or at −86°C until gel electrophoresis can be performed. SDS-PAGE of Soluble Whole-Cell Proteins. For protein electrophoresis 15 µL of the centrifuged supernatant from each strain are applied directly onto the surface of an 0.5 mm thin precast polyacrylamide 8%–18% (w/v) gradient gel (ExcelGel SDS, Amersham Pharmacia). The proteins are separated using a horizontal Multiphor II electrophoresis apparatus with a power supply (MultiDrive XL) at 600  V, 50 mA, and 30 W at a constant cooling temperature of 15°C (MultTemp II) for 90 min following the manufacturer’s instructions. A molecular weight protein ladder standard is included in each electrophoretic run. Each strain is electrophoresed in triplicate, involving the entire experimental procedure from culture to extraction. After electrophoresis, the SDS gel is examined with silver nitrate under constant shaking. The gels are fixed overnight in fixative (50% methanol, 37% formaldehyde) solution, followed by treatment in dithiothreitol (DTT) solution (1 mL DTT stock) (100 mg DTT/20 mL ddH2O)/1000 mL for 45 min. The gels are stained with silver nitrate for 60 min and washed for 30 s in deionized distilled water. Thereafter, the gels are washed twice with 100 mL developer solution (0.28 M sodium carbonate, 37% formaldehyde solution), each time for 30 s, and suspended in developer solution for additional 5 min or until the bands are viable. To prevent the gel from overstaining, citric acid is added directly to the developer solution and allowed to settle down for 10 min. Finally, the gel is washed with deionized distilled water, photographed, and stored. Band Pattern Analysis. The strain differences in band patterns generated by SDS-PAGE are first established visually by recording the presence or absence of bands without considering intensity. SDS-PAGE electrophoresis bands on gels from different isolates are compared using the Dendron (Solltech Inc,) computer-assisted program. If the similarity coefficients (SAB) for the patterns of two strains are identical, the SAB is equal to 1.00. If one or more bands differ in size and intensity, there is a gradation of SAB from 0.99 to 0.01, with decreasing similarity. An SAB of 0.00 indicates that no bands in two patterns are of the same size. Dendrograms based on SAB are generated using the Dendron computer program with the unweighted pair group method. In dendrogram construction, the program first searches for the two strains with the highest SAB and groups them into a unit with a branching point corresponding to their SAB. The program then searches again for the strain-strain or strain-unit pair with the highest SAB and connects it. The process continues until all isolates are connected. A branching point connecting a unit and a strain is determined by the average SAB between the member of the unit and another strain or unit.

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(ii) Extraction of DNA for Genetic Analysis  For genetic detection and analysis, DNA is extracted using the guanidium procedure.5,6 Leptotrichia pellets are resuspended in TE buffer (10 mM Tris-HCl–1 mM EDTA, pH 8.0) and incubated with 10% SDS, lysozyme, and proteinase K (20 mg/ mL) for 30 min at 37°C. The cells are lyzed with GES reagent (5  M guanidium thiocyanate, 100 mM EDTA and 0.5% sarkosyl). After cooling the sample on ice for 5 min, 7.5 M cold ammonium acetate is added. The phases are mixed gently by shaking and are kept on ice for additional 10 min. Chloroform-2-pentanol (24:1) is then added. Again, the phases are mixed by vigorous shaking, and centrifuged for 5 min at 13,000 × g. The aqueous phases are carefully transferred to a 1.5 mL Eppendorf tube. This procedure is repeated. Cold isopropanol (0.54 volumes) is added to a new tube containing the aqueous phase. The tube is inverted until the DNA precipitates, and centrifugation occurs at 13,000 × g for 4 min. The aqueous solution is aspirated, and the pellet is washed with 70% ethanol. A vacuum concentrator is used to dry the DNA for 15 min. The pellet is dissolved in TE buffer and treated with DNase-free RNase for 1 h at 37°C. Chloroform2-pentanol is added and mixed by vigorous shaking, followed by centrifugation for 5 min at 13,000 × g. The aqueous phase is overlaid with 0.54 volume of cooled isopropanol (−20°C). The tube is inverted until the DNA precipitates and centrifugation occurs at 13,000 × g for 4 min. The aqueous solution is discarded, and the pellet washed with 70% ethanol. A vacuum concentrator is used to dry the DNA for 15 min. The DNA pellet is dissolved in TE buffer or deionized distilled water and stored at 4°C. To check the quality of the extracted DNA, a total volume of 6 µL is made up by mixing 1 µL of DNA, 3 µL of 10 × loading buffer plus ddH2O. The mixture is loaded into 0.7% agarose gel and electrophoresed for 1 h at 150 V using a horizontal gel electrophoresis apparatus. The gel is stained with ethidium bromide for 30 min. The stain is removed with ddH2O, and the gel is inspected under UV illumination. Only one band of chromosomal DNA is observed. No smears are seen. To insure good quality DNA for RAPD analysis, each DNA preparation is measured at ODs 260/280 and the absorbance ratio is recorded. Samples with ratios below or above 1.8–1.9 are reextracted.

49.2.2  Detection Procedures RAPD Analysis. Amplification reactions for Leptotrichia are performed in GeneAmp PCR reaction tubes,6 each containing a final volume of 20 µL, consisting of (i) 11.2 µL sterile double-distilled water; (ii) 1× buffer [10× PCR buffer (10 mM Tris-HCl, pH 8.3, 50 mM KCl)]; (iii) 1 mM dNTPs (10 mM); (iv) 3U (10 U/µL AmpliTaq DNA Polymerase Stoffel fragment (5 U/µL); (v) 2 µM primer (20 µM) (Operon); (vi) 1.5 mM MgCl2 (25 mM); and (vii) 1.0 µL of 1:5 diluted DNA template (1–10 ng/µL). A master mix of the above reaction mixture is made in the order (i–vi) mixed together, and aliquots are made into individual MicroAmp reaction tubes. Component (vii) is added directly into each tube. All these

Molecular Detection of Human Bacterial Pathogens

steps are performed on ice. A negative control without DNA is included in each experiment together with a DNA molecular weight marker: 1 kb Ladder Plus. DNA is amplified in a Perkin Elmer GeneAmp PCR System 2400. The thermal cycler temperature profile consists of denaturation at 96°C for 5 min, followed by 35 cycles of 96°C for 1 min, 48°C for 5 min, and 74°C for 1 min. Five µL of amplification products are analyzed by electrophoresis in 1.5% agarose gel on a horizontal gel electrophoresis apparatus. Gels are stained with ethidium bromide, and bands are visualized and photographed by using a UV transilluminator and a Kodak digital scienceTM electrophoresis gel documentation system 120. Amplification of 16S rRNA and Purification of PCR Products. The 16S rRNA genes from the examined Leptotrichia strains are amplified under standardized conditions with the primers 9F and 1541R.5 PCR is performed in thin-walled tubes with a Perkin-Elmer 9700 thermo cycler. One microliter of the diluted (1:5) DNA template is added to a reaction mixture (50 µL final volume) containing 1× Taq 2000 reaction buffer, 2.5 mM MgCl2, 0.8 µM dNTPs, 400 nM of each primer, and 1 U Taq 2000 polymerase (Stratagene) in buffer containing Taqstart antibody (Sigma). In a hot-start protocol, samples are preheated at 94°C for 4 min, followed by amplification with 30 cycles of 94°C for 45 s, 60°C for 45 s, and 72°C for 1.5 min, with an additional 1 s for each cycle; and a final step at 72°C for 15 min. The PCR products are examined by electrophoresis in a 1% agarose gel. DNA is stained with ethidium bromide and visualized under short-wavelength UV light. Amplified 16S rRNA genes are purified with the Sequenase kit for partial sequencing and on Sephadex G-50 resin columns before full-length sequencing. 16S rRNA Sequencing. Purified PCR is sequenced using an ABI Prism cycle-sequencing kit (BigDye Terminator cycle-sequencing kit). Quarter-dye chemistry is applied with 3.2 µM primers and 1.5 µL PCR products in a final volume of 20 µL. Cycle sequencing is done with an ABI 9700 PCR machine with 25 cycles of denaturation at 96°C for 10 s and annealing and extension at 60°C for 4 min. Purified DNA products are dried in a speed vac centrifuge for 75 min, resuspended in 3.2 µL loading dye, and denatured at 98°C for 2 min. Sequencing reactions are run on an ABI Prism 377 DNA sequencer. Data Analyses of 16S rRNA Sequences. To determine the identity or approximate phylogenetic position of the Leptotrichia strains,5 410 bases are first sequenced. Using six additional sequencing primers,5 full sequences (about 1500 bases) are obtained for representative strains. For identification of the closest relatives, the sequences of the unrecognized Leptotrichia strains are compared with the 16S rRNA gene sequences of over 9000 bacteria in Paster and Dewhirst’s database and 76,000 sequences in the Ribosomal Data Project.105 Similarity matrices are corrected for multiple base changes at single positions by the method of Jukes and Cantor.106 Phylogenetic trees are constructed by the neighborjoining method of Saitou and Nei.107 Sequences are aligned using the MegAlign program (DNASTAR) and imported into

Leptotrichia and Leptotrichia-Like Organisms

TREECON, a software package for the Microsoft Windows environment, which is used for the construction and drawing of evolutionary trees.108 DNA–DNA Relatedness Assays. The S1-nuclease procedure for the free-solution reassociation for DNA similarity assays is used for pair wise reactions5,109 with selected strains of Leptotrichia. All procedures, including DNA isolation, French pressure cell fragmentation, hybridization, and S1-nuclease assays have been detailed elsewhere.109 However, rather than labeling the probe DNA chemically with 125I, the random primers method is used (Random Primers DNA Labeling System) to label DNA with (α-33P)dCTP (Perkin Elmer Life Sciences). The probe and target DNAs are reassociated at 66.2°C ± 0.5°C for 24 h for Leptotrichia DNA, with a mean G + C content of 29.7 mol%.55 Values for both homologous and heterologous reassociations are corrected for nonspecific heteroduplex formation by control reactions with salmon sperm single-stranded DNA (0% DNA relatedness) and are less than 10%. Each reaction is repeated at least three times. The mean of all reactions is reported as percentage DNA–DNA relatedness. DNA from Erwinia amylovora ATCC 29780 (EA 110) is used as outgroup.

49.3  C  ONCLUSION AND FUTURE PERSPECTIVES There is no doubt that Leptotrichia are agents of emerging infections, although they are considered members of the normal flora in the oral cavity and genital tract. A particular reason for this is that improvements in medical treatment make people live longer. This causes a number of compromised hosts that are susceptible to indigenous Leptotrichia, particularly after breakage of mucosal barriers. Such breakage may serve as a portal of entry for Leptotrichia. However, Leptotrichia can also cause infections of the noncompromised host. This makes Leptotrichia more than opportunists. With such a wide pathogenic potential Leptotrichia infection should be expected relatively frequently. Up to now, this has not been so. A reason for that is that Leptotrichia can easily be confused with other bacteria, such as Lactobacillus, Fusobacterium, and Capnocytophaga. Furthermore, Leptotrichia infections cannot be said to have distinct clinical characteristics, although certain reactions such as fever may be ascribed to their lipopolysaccharide. The isolation of Leptotrichia can be a challenge to both microbiologist and clinician. Recognition of a characteristic morphology together with a typical carbohydrate fermentation pattern with lactic acid as major end product after culture in PYG broth in an appropriate clinical setting should raise the suspicion of Leptotrichia infection. Leptotrichia are often considered contaminants if isolated from clinical infection, which may be unfair, particularly if the host is compromised. Recovery of Leptotrichia requires adherence to strict anaerobic techniques during clinical sampling, transport, and culture. Handling in the microbiology laboratory should also be completely anaerobic although a few strains may be aerotolerant and become used to aerobicity during

563

repeated culture. Adequate experience in microbial diagnostics, taxonomy, and equipment are required to bring forward a correct Leptotrichia diagnosis. Unfortunately, not all laboratories are equipped for this purpose. The taxonomic situation of Leptotrichia has been far from clear. Relatively few Leptotrichia species have been properly established. Species such as “L. amnionii” is still pending valid publication. A number of Leptotrichia strains have been listed in databases but cannot be cultured. We think that more efforts should be made to culture not-yet-cultivable Leptotrichia. This is important with regard to learning more about their pathogenic role. We also think that methods should be developed to test the possible significance of not-yet-cultivated strains in clinical settings. Little is known about the pathogenicity of Leptotrichia. Aside from a few studies on the toxicity of lipopolysaccharide, which is high compared to that of other organisms in the oral flora, practically nothing has been done. Leptotrichia often occurs together with other species. We do not know about synergistic mechanisms, although they may play a role. Nor do we know much about the host interactions. Little is also known of their immunological reactions, although antibodies are raised to the lipopolysaccharide of Leptotrichia. Because Leptotrichia can cause emerging infections, we should indeed pay more attention to the clinical significance and microbiology of these bacteria.

ACKNOWLEDGMENTS This work was supported in part by the Faculty of Dentistry, University of Oslo and the Norwegian Geotechnical Institute, Norway. The support is highly appreciated and acknowledged.

REFERENCES







1. Trevisan, V., Prime linee d´introduzione allo studio dei Batterj italiani. Rendiconti dell´instituto Lombardo, di Scienze et letter. Series 2, 12, 133, 1879. 2. Hamilton, R.D., and Zahler, S.A., A study of Leptotrichia buccalis, J. Bacteriol., 73, 386, 1957. 3. Eribe, E.R.K., and Olsen, I., Leptotrichia species in human infections, Anaerobe, 14, 131, 2008. 4. Ludwig, W., Euzéby, J., and Whitman, W.B., Draft taxonomic outline of the Bacteroidetes, Planctomycetes, Chlamydiae, Spirochaetes, Fibrobacteres, Fusobacteria, Acidobacteria, Verrucomicrobia, Dictyoglomi, and Gemmatimonadetes, www.bergeys.org/outlines/bergeys_vol_4-outline.pdf, last acc­ essed November 3, 2010. 5. Eribe, E.R.K. et al., Genetic diversity of Leptotrichia and description of Leptotrichia goodfellowii sp. nov., Leptotrichia hofstadii sp. nov., Leptotrichia shahii sp. nov., and Leptotrichia wadei sp. nov., Int. J. Syst. Evol. Microbiol., 54, 583, 2004. 6. Eribe, E.R., and Olsen, I., SDS-PAGE of whole-cell proteins and random amplified polymorphic DNA (RAPD) analyses of Leptotrichia isolates, Microbial Ecol. Health Dis., 14, 193, 2002. 7. Paster, B.J. et al., Prevalent bacterial species and novel phylotypes in advanced noma lesions, J. Clin. Microbiol., 40, 2187, 2002.

564









8. Marques da Silva, R. et al., Bacterial diversity in aortic aneurysms determined by 16S ribosomal RNA gene analysis, J. Vasc. Surg., 44, 1055, 2006. 9. Goldstein, E.C. et al., Bacteriology of human and animal bite wounds, J. Clin. Microbiol., 8, 667, 1978. 10. Hirsch, D.C., Biberstein, E.L., and Jang, S.S., Obligate anaerobes in clinical veterinary practice, J. Clin. Microbiol., 10, 188, 1979. 11. Tindall, B.J., Taxonomic note. Rule 15 of the international code of nomenclature of bacteria: A current source of confusion, Int. J. Syst. Evol. Microbiol., 58, 1775, 2008. 12. Tee, W. et al., Bacteremia due to Leptotrichia trevisanii sp. nov., Eur. J. Clin. Microbiol. Infect. Dis., 20, 765, 2001. 13. Shukla, S.K. et al., Leptotrichia amnionii sp. nov., a novel bacterium isolated from the amniotic fluid of a woman after intrauterine fatal demise, J. Clin. Microbiol., 40, 3346, 2002. 14. Page, L.R., and Krywolap, G.N., Determination of the deoxyribonucleic acid composition and deoxyribonucleic acid-­deoxyribonucleic acid hybridization of Fusobacterium fusiforme, Fusobacterium polymorphum, and Leptotrichia buccalis: taxonomic considerations, Int. J. Syst. Bacteriol., 26, 301, 1976. 15. Hofstad, T., and Olsen, I., Fusobacterium and Leptotrichia. In Borrelier, S.P. (Editor), Topley and Wilson’s Microbiology and Microbial Infections, Bacteriology, 10th ed., vol. 2, p. 1945, Arnold, London, 2005. 16. Bennett, K.W., and Duerden, B.I., Identification of fusobacteria in a routine diagnostic laboratory, J. Appl. Bacteriol., 59, 171, 1985. 17. Hofstad, T., and Selvig, K.A., Ultrastructure of Leptotrichia buccalis, J. Gen. Microbiol., 56, 23, 1969. 18. Listgarten, M.A., and Lai, C.H., Unusual cell wall ultrastructure of Leptotrichia buccalis, J. Bacteriol., 123, 747, 1975. 19. Smith, S.H., Murray, R.G., and Hall, M., The surface structure of Leptotrichia buccalis, Can. J. Microbiol., 40, 90, 1994. 20. Könönen, E. et al., The oral gram-negative anaerobic microflora in young children: Longitudinal changes from edentulous to dentate mouth, Oral Microbiol. Immunol., 9, 136, 1994. 21. George, W.L. et al., Gram-negative anaerobic bacilli: Their role in infection and patterns of susceptibility to antimicrobial agents. II. Little-known Fusobacterium species and miscellaneous genera, Rev. Infect. Dis., 3, 599, 1981. 22. Söderberg, G., Lindberg, A.A., and Nord, C.E., Bacteroides fragilis in acute salpingitis, Infection, 7, 226, 1979. 23. Newman, M.G., et al., Predominant microbiota associated with periodontal health in the aged, J. Periodontol., 49, 553, 1978. 24. Russell, C., and Melville, T.H., Bacteria in the human mouth, J. Appl. Bacteriol., 44, 163, 1978. 25. Holdeman, L.V., Cato, E.P., and Moore, W.E.C., Anaerobe Laboratory Manual, 4th ed., Virginia Polytechnic Institute and State University, Blacksburg, 1977. 26. Moore, W.E.C., Johnson, J.L., and Holdeman, L.V., Emendation of Bacteroidaceae and Butyrivibrio and descriptions of Desulfomonas gen. nov. and ten new species in the genera Desulfomonas, Butyrivibrio, Eubacterium, Clostridium and Ruminococcus, Int. J. Syst. Bacteriol., 26, 238, 1976. 27. Williams, B.L., Pantalone, R.M., and Sherris, J.C., Subgingival microflora and periodontitis, J. Peridontol. Res., 11, 1, 1976. 28. Drasar, B.S., and Hill, M.J., Human Intestinal Flora, Academic Press, London, 1974, 263. 29. Hardie, J.M., and Bowden, G.H., The normal microbial flora of the mouth. In Skinner, F.A., and Carr, J.G. (Editors), The Normal Microbial Flora of Man, p. 47, Academic Press, London, 1974.

Molecular Detection of Human Bacterial Pathogens 30. Evaldson, G. et al., Microbiological findings in pregnant women with premature rupture of the membranes, Med. Microbiol. Immunol., 168, 283, 1980. 31. Könönen, E., Asikainen, S., and Jousimes-Somer, H., The early colonization of gram-negative anaerobic bacteria in edentulous infants, Oral Microbiol. Immunol., 7, 28, 1992. 32. Rosebury, T., Microorganisms Indigenous to Man, p. 435, McGraw-Hill, New York, 1962. 33. Colombo, A.P. et al., Clinical and microbiological features of refractory periodontitis subjects, J. Clin. Periodontol., 25, 169, 1998. 34. Duperval, R., Béland, S., and Marcoux, J.A., Infective endocarditis due to Leptotrichia buccalis: A case report, Can. Med. Assoc. J., 130, 422, 1984. 35. Kohler, J.L. et al., Septicémie a Leptotrichia buccalis chez un immunodéprimé, Pathol. Biol., 30, 161, 1982. 36. Morgenstein, A.A. et al., Serious infection with Leptotrichia buccalis. Report of a case and review of the literature, Am. J. Med., 69, 782, 1980. 37. Thadepalli, H., and Rao, B., Clinical evaluation of mezlocillin, Antimicrob. Agents Chemother., 16, 605, 1979. 38. Krywolap, C.N., and Page, L.R., Oral Fusobacterium, Leptotrichia and Bacterionema: II. Pathogenicity: A review of the literature, J. Baltimore Coll. Dent. Surg., 32, 26, 1977. 39. Gupta, H., and Matlow, A., Leptotrichia buccalis causing cellulitis and bacteremia in a neutropenic child: A case report, Clin. Microbiol. Newslett., 20, 195, 1998. 40. Benson, P.M., Sexually transmitted diseases, in Military Dermatol. http://www.heart-intl.net/HEART/110105/Sexually Transmitted.pdf. 41. Hebb, J.K. et al., Detection of novel organisms associated with salpingitis, by use of 16S rDNA polymerase chain reaction, J. Infect. Dis., 190, 2109, 2004. 42. Pieper, R. et al., Acute appendicitis and Bacteroides fragilis, Scand. J. Infect. Dis. Suppl., 19, 92, 1979. 43. Fredricks, D.N., Schubert, M.M., and Myerson, D., Molecular identification of an invasive gingival bacterial community, Clin. Infect. Dis., 41, e1, 2005. 44. Moore, W.E.C., et al., Bacteriology of experimental gingivitis in children, Infect. Immun., 46, 1, 1984. 45. Nakou, M. et al., Early microbial colonization of permucosal implants in edentulous patients, J. Dent. Res., 66, 1654, 1987. 46. Vernelen, K. et al., Bacteraemia with Leptotrichia buccalis: Report of a case and review of the literature, Acta Clin. Belg., 51, 265, 1996. 47. van Houte, J., Role of micro-organisms in caries etiology, J. Dent. Res., 73, 672, 1994. 48. Hofstad, T., Pathogenicity of anaerobic gram-negative rods: Possible mechanisms, Rev. Infect. Dis., 6, 189, 1984. 49. Preza, D. et al., Bacterial profiles of root caries in elderly patients, J. Clin. Microbiol., 46, 2015, 2008. 50. Grollier, G. et al., Leptotrichia buccalis isolated from peritoneal fluid of two immunocompromised patients, Clin. Microbiol. Newslett., 12, 62, 1990. 51. Caram, L.B. et al., Leptotrichia endocarditis: Report of two cases from the International Collaboration on Endocarditis (ICE) database and review of previous cases, Eur. J. Clin. Microbiol. Infect. Dis., 27, 139, 2008. 52. Lark, R.L. et al., Risk factors for anaerobic bloodstream infections in bone marrow transplant recipients, J. Clin. Infect. Dis., 33, 338, 2001. 53. Patel, J.B. et al., Bacteremia caused by a novel isolate resembling Leptotrichia species in a neutropenic patient, J. Clin. Microbiol., 37, 2064, 1999.

Leptotrichia and Leptotrichia-Like Organisms 54. Messiaen, T., Lefebvre, C., and Geubel, A., Hepatic abscess likely related to Leptotrichia buccalis in an immunocompetent patient, Liver, 16, 342, 1996. 55. Schwartz, D.N. et al., Leptotrichia buccalis bacteremia in patients treated in a single bone marrow transplant unit, Clin. Infect. Dis., 20, 762, 1995. 56. Hamman, R. et al., Endocarditis caused by a Leptotrichia buccalis-like bacterium in a patient with a prosthetic aortic valve, Eur. J. Clin. Microbiol. Infect. Dis., 12, 280, 1993. 57. Weinberger, M. et al., Leptotrichia buccalis bacteremia in patients with cancer: Report of four cases and review, Rev. Infect. Dis., 13, 201, 1991. 58. Baquero, F. et al., Capnophilic and anaerobic bacteria in a neutropenic patient: An oral source, Rev. Infect. Dis., 12, 157, 1990. 59. Reig, M. et al., Leptotrichia buccalis bacteremia in neutropenic children, J. Clin. Microbiol., 22, 320, 1985. 60. Prévot, A.R., Septicémie a “Leptotrichia vaginalis,” Sem. Hop., 30, 2040, 1968. 61. Sutter, V.L., Anaerobes as normal oral flora, Rev. Infect. Dis., 6, Suppl 1, S62, 1984. 62. Robin, C., Histoire Naturelle des Végétaux Parasites qui Croissent sur l’Homme et sur les Animaux vivants, XVI, Baillière, Paris, 1853, 702. 63. Wherry, W.B., and Oliver, W.W., Leptothrix innominata (Miller), J. Infect. Dis., 19, 299, 1916. 64. Thjøtta, T., Hartmann, O., and Bøe, J., A study of the Leptotrichia trevisan. History, morphology, biological and serological characteristics. Skr. Nor. Vidensk.- Akad. Oslo. I. Mat.-Naturvidensk. Kl., 5, 1, 1939. 65. Böe, J., and Thjötta, T., The position of Fusobacterium and Leptotrichia in the bacteriological system, Acta Pathol. Microbiol. Scand., 21, 441, 1944. 66. Gmür, R., et al., Gingival crevice microbiota from Chinese patients with gingivitis or necrotizing ulcerative gingivitis, Eur. J. Oral. Sci., 112, 33, 2004. 67. Ruviére, D.B. et al., Assessment of the microbiota in root canals of human primary teeth by checkerboard DNA–DNA hybridization, J. Dent. Child. (Chic), 74, 118, 2007. 68. Sassone. L.M. et al., Microbiological evaluation of primary endodontic infections in teeth with and without sinus tract, Int. Endod. J., 41, 508, 2008. 69. Sassone, L. et al., Evaluation of the microbiota of primary endodontic infections using checkerboard DNA–DNA hybridization, Oral Microbiol. Immunol., 22, 390, 2007. 70. Tyrrell, K.L. et al., Anaerobic bacteria cultured from the tongue dorsum of subjects with malodour, Anaerobe, 9, 243, 2003. 71. Fredricks, D.N. et al., Changes in vaginal bacterial concentrations with metronidazole therapy for bacterial vaginosis as assessed by quantitative PCR, J. Clin. Microbiol., 2009, January 14. (Epub ahead of print). 72. Fredricks, D.N. et al., Targeted PCR for detection of vaginal bacteria associated with bacterial vaginosis, J. Clin. Microbiol., 45, 3270, 2007. 73. Tamrakar, R. et al., Association between Lactobacillus species and bacterial vaginosis-related bacteria, and bacterial vaginosis scores in pregnant Japanese women, BMC Infect. Dis., 7, 128, 2007. 74. Fredricks, D.N., Fiedler, T.L., and Marazzo, J.M., Molecular identification of bacteria associated with bacterial vaginosis, N. Engl. J. Med., 353, 1899, 2005. 75. Fredricks D.N., and Marrazzo, J.M., Molceular methodology in determining vaginal flora in health and disease: Its time has come, Curr. Infect. Dis. Rep., 7, 463, 2005.

565 76. Thilesen, C.M. et al., Leptotrichia amnionii, an emerging pathogen of the female urogenital tract, J. Clin. Microbiol., 45, 2347, 2007. 77. Boennelycke, M. et al., Leptotrichia amnionii found in septic abortion in Denmark, Scand. J. Infect. Dis., 39, 382, 2007. 78. Domann, E. et al., Culture-independent identification of pathogenic bacteria and polymicrobial infections in the genitourinary tract of renal transplant recipients, J. Clin. Microbiol., 41, 5500, 2003. 79. DiGiulio, D.B. et al., Microbial prevalence, diversity and abundance in amniotic fluid during preterm labor: A molecular and culture-based investigation, PLoS ONE, 3, e 3056, 2008. 80. Han, Y.W. et al., Uncultivable bacteria as etiologic agents of intra-amniotic inflammation leading to preterm birth, J. Clin. Microbiol., 47, 38, 2009. 81. Goto, M., Hitomi, S., and Ishii, T., Bacterial arthritis caused by Leptotrichia amnionii, J. Clin. Microbiol., 45, 2082, 2007. 82. Hot, A. et al., Lemierre syndrome caused by Leptotrichia buccalis in a neutropenic patient, Int. J. Infect. Dis., 12, 339, 2008. 83. Bhally, H.S. et al., Leptotrichia buccalis bacteremia in two patients with acute myelogenous leukemia, Anaerobe, 11, 350, 2005. 84. De Martino, S.J. et al., Peripartum bacteremias due to Leptotrichia amnionii and Sneathia sanguinegens, rare causes of fever during and after delivery, J. Clin. Microbiol., 42, 5940, 2004. 85. Ulstrup, A.K., and Hartzen, S.H., Leptotrichia buccalis: A rare cause of bacteraemia in non-neutropenic patients, Scand. J. Infect. Dis., 38, 712, 2006. 86. Falkler, W.A., Jr., Hawley, C.E., and Mongiello, J.R., Leptotrichia buccalis hemagglutination in cell binding and salivary inhibition studies, J. Periodontal Res., 13, 425, 1978. 87. Kondo, W., Sato, M., and Ozawa, H., Haemagglutinating activity of Leptotrichia buccalis cells and their adherence to saliva-coated enamel powder, Arch. Oral Biol., 21, 263, 1976. 88. Gustafson, R.L. et al., The biological activity of Leptotrichia buccalis endotoxin, Arch. Oral Biol., 1, 1149, 1966. 89. Dearaujo, W.C., Varah, E., and Mergenhagen, S.E., Immunochemical analysis of human oral strains of Fusobacterium and Leptotrichia, J. Bacteriol., 86, 837, 1963. 90. Patters, M.R. et al., Blastogenic response of human lymphocytes to oral bacterial antigens: Comparison of individuals with periodontal disease to normal and edentulous subjects, Infect. Immun., 14, 1213, 1976. 91. Schüpbach, P., Osterwalder, V., and Guggenheim, B., Human root caries: Microbiota in plaque covering sound, carious and arrested caries root surfaces, Caries Res., 29, 382, 1995. 92. Hooper, S.J. et al., A molecular analysis of the bacteria present within oral squamous cell carcinoma, J. Med. Microbiol., 56, 1651, 2007. 93. McNamara, T.F. et al., The development of an altered gingival crevicular microflora in the alloxan-diabetic rat, Arch. Oral Biol., 27, 217, 1982. 94. Kawanami, T. et al., Severe pneumonia with Leptotrichia sp. detected predominantly in bronchoalveolar lavage fluid by use of 16S rRNA gene sequencing analysis, J. Clin. Microbiol., 47, 498, 2009. 95. Haggerty, C.L. et al., Clinical characteristics of bacterial vaginosis among women testing positive for fastidious bacteria, Sex. Transm. Infect., 2008 November 12. (Epub ahead of print).

566 96. Gundi, V.A.K.B., Desbriere, R., and La Sola, B., Leptotrichia amnionii and the female reproductive tract, Emerg. Infect. Dis., 10, 2056, 2004. 97. Eribe, E.R., Hofstad, T., and Olsen, I., Enzymatic/biochemical and cellular fatty acid analyses of Leptotrichia isolates, Microbial Ecol. Health Dis., 14, 137, 2002. 98. Hofstad, T., Fusobacterium and Leptotrichia. In Collier, L., Balows, A., and Sussman, M. (Editors), Topley and Wilson’s Microbiology and Microbial Infections, 9th ed., vol. 3,, 1356, Arnold, London, 1998. 99. Shah, H.N. et al., Approaches to the study of the systematics of anaerobic, gram negative, non spore-forming rods; current status and perspectives, Anaerobe, 2009, doi:10.1016/j. anaerobe.2009.08.003. 100. Olsen, I., and Shah, H.N., International Committee on Systematics of Prokaryotes. Subcommittee on the taxonomy of Gram-negative anaerobic rods. Minutes of the open and closed meeting, 2–3 September 2007, Edinburgh, UK, Int. J. Syst. Evol. Microbiol., 58, 1051, 2008. 101. Murray, P.R., and Citron, D.M., Anaerobic Gram-negative bacilli and cocci. In Balows, A. (Editor), Manual of Clinical Microbiology, 5th ed., p. 488, ASM, Washington, DC, 1991. 102. McCarthy, C., and Snyder, M.L., Selective medium for fusobacteria and leptotrichia, J. Bacteriol., 86, 158, 1963.

Molecular Detection of Human Bacterial Pathogens 103. Bernard, K. et al., Use of chemotaxonomy as an aid to differentiate among Capnocytophaga species, CDC group DF-3, and aerotolerant strains of Leptotrichia buccalis, J. Clin. Microbiol., 29, 2263, 1991. 104. Olsen, I. et al., Cultivated and not-yet-cultivated bacteria in oral biofilms, Microbial Ecol. Health Dis., 21, 1, 2009. 105. Cole, J.R., et al., The Ribosomal Database Project (RDP-II): Previewing a new autoaligner that allows regular updates and the new prokaryotic taxonomy, Nucleic Acids Res., 31, 442, 2003. 106. Jukes, T.H., and Cantor, C.R., Evolution of protein molecules. In Munro, H.N. (Editor), Mammalian Protein Metabolism, p. 21, Academic Press, New York, 1969. 107. Saitou, N., and Nei, M., The neighbor-joining method: A new method for reconstructing phylogenetic trees, Mol. Biol. Evol., 4, 406, 1987. 108. Van de Peer, Y. et al., Database on the structure of small ribosomal subunit RNA, Nucleic Acids Res., 24, 86, 1996. 109. Johnson, J.L., Similarity analysis of DNAs. In Gerhardt, P., Murray, R. G.E., Wood, W.A., and Krieg, N.R. (Editors), Methods for General and Molecular Bacteriology, p. 655, American Society for Microbiology, Washington, DC, 1994.

50 Porphyromonas Stefan Rupf, Wolfgang Pfister, and Klaus Eschrich CONTENTS 50.1 Introduction...................................................................................................................................................................... 567 50.1.1 Classification, Morphology, and Epidemiology......................................................................................... 567 50.1.2 Pathogenesis and Clinical Features............................................................................................................ 569 50.1.3 Diagnostics......................................................................................................................................................... 573 50.1.3.1 Conventional Techniques....................................................................................................................... 573 50.1.3.2 Molecular Techniques............................................................................................................................ 573 50.2 Methods............................................................................................................................................................................ 574 50.2.1 Sample Preparation.......................................................................................................................................... 574 50.2.2 Detection Procedures...................................................................................................................................... 575 50.2.2.1 Species-Specific PCR............................................................................................................................ 575 50.2.2.2 Quantitative Real-Time PCR................................................................................................................. 575 50.2.2.3 Quantitative Competitive PCR (QC–PCR)............................................................................................ 576 50.3 Conclusions and Perspectives........................................................................................................................................... 576 References.................................................................................................................................................................................. 577

50.1  INTRODUCTION 50.1.1  Classification, Morphology, and Epidemiology Classification. The genus Porphyromonas is classified to the family of Bacteroidaceae. So far, 17 species have been described (Table 50.1). Although Porphyromonas is closely related phylogenetically to the genera Bacteroides and Prevotella, Shah and Collins suggested a new taxon after reclassification of the Bacteroides spp. B. asaccharolyticus, B. gingivalis, and B. endodontalis on the basis of their biochemical and chemical properties.1,3,6,27–29 Then 16S rRNA gene sequencing confirmed the independent taxon Porphyromonas. The phylogenetic differentiation of the members of the genus Porphyromonas was also evidenced by 16S rRNA gene sequence analysis and confirmed by DNA–DNA relatedness studies and by morphological, chemical, and biochemical criteria.30–32 The discrimination of the species P. gingivalis and P. gulae was enabled by comparing 16S-23S rRNA gene internal transcribed spacer sequences.8,21 Members of the Porphyromonas genus have been isolated from a number of mammals. The species P. asaccharolytica,1 P. bennonis,4 P. catoniae,5 P. endodontalis,1 P. gingivalis,1 P. somerae,7 and P. uenonis 9 have been found worldwide in humans. Several Porphyromonas species, P.  macacae,26 P. cansulci,10 P. canoris,12 P. cangingivalis,10 and P. circumdentaria,15 were isolated from animals, where they are mostly associated

with oral inflammatory processes. These species were frequently found in humans in infected dog or cat bite wounds.33 Porphyromonas spp. are generally associated with inflammation and purulence, but some species primarily occur in gingival and periodontal disease processes, occasionally then causing accompanying systemic inflammative processes. P. gingivalis is closely associated with the progression of chronic periodontitis. The genus Porphyromonas is subject to modifications. A number of strains isolated from human oral samples are genetically heterogeneous with regard to their 16S rRNA genes. These strains—among them Porphyromonas P334 — have not yet been comprehensively characterized. Numerous other strains primarily of oral origin are listed on the species level in the NCBI taxonomy database.35,36 P. macacae used to be referred to as P. salivosa.15 A species that was originally described as “Porphyromonas denticanis”37 was reclassified as Odoribacter denticanis. Morphology. The members of the genus Porphyromonas have been described as strictly anaerobic, nonmotile, nonspore-forming, gram-negative rods or coccoid-shaped. Depending on their stage of growth, the size of the pleomorphic Porphyromonas differs from 0.4 to 0.8 by 0.8–6 µm. They are equipped with the enzymes glutamate dehydrogenase and superoxide dismutase, but lack glucose-6-­phosphatedehydrogenase and 6-phosphogluconatedehydrogenase. Their metabolism is proteolytic. Energy is produced by fermentation of amino acids. Carbohydrates are not fermented. The optimal pH of 567

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TABLE 50.1 The Genus Porphyromonas: Classification, Origin, and Association with Diseases of the Members of the Genus (in Alphabetic Order) Species Human Origin P. asaccharolytica1 (B. melaninogenicus subsp. asaccharolyticus2; B. asaccharolyticus3) P. bennonis4 P. catoniae (Oribaculum catoniae5) P. endodontalis1 (B. endodontalis)6

P. gingivalis1

P. somerae (P. levii-like organisms)7 P. uenonis (P. endodontalis-like organisms)8,9 Animal Origin P. asaccharolytica1 P. cangingivalis10,11 P. canoris12,13 P. cansulci10 P. catoniae5,13,14 P. circumdentaria15–17 P. crevioricanis11,19,20 P. endodontalis16 P. gingivalis11,21,22,23 (differentiation) into P. gulae (animal origin) and P. gingivalis (human origin) P. gingivicanis11,19,20 P. gulae23 P. levii7,24,25 P. macacae (Syn: P. salivosa)26

Source, Associated with

Cultural and Biochemical Characteristics

Abscesses from different regions, nonoral P: +; I: +; C: −; Gf: −; α-F: +; β-NAG − mucosal surfaces, periodontal pocket Wound infections and abscesses Healthy and diseased human gingival, shallow pockets Apical periodontitis, apical root canal, periodontitis, periodontal pocket, carious dentine Periodontitis, oral cavity, periodontal pocket, apical periodontitis, root canal, carious dentine Skin and soft-tissue abscesses Gastrointestinal tract

P: weakly +; I: −; C: different; Gf: −; α-F: −; β-NAG + P: −; I: −; C: −; Gf: +; α-F: +; β-NAG +

Footrot, goat Dental plaque, periodontal pocket, dog Plaque, periodontal pocket, dog Plaque, periodontal pocket, dog Gastrointestinal tract, piglet Plaque, gingival margins, dog, cats Plaque, dog Plaque, dog Plaque, monkey, dog

P: +; I: +; C: −; Gf: −; α-F: +; β-NAG − P: +; I: +; C: +; Gf: most strains −; α-F: −; β-NAG different P: +; I: +; C: +; Gf: most strains −; α-F: −; β-NAG different P: +; I: +; C: +; Gf: most strains −; α-F: −; β-NAG different P: −; I: −; C: −; Gf: +; α-F: +; β-NAG + P: +; I: +; C: +; Gf: most strains −; α-F: −; β-NAG different P: +; I: +; C: +; Gf: most strains −; α-F: −; β-NAG different P: +; I: +; C: −; Gf: −; α-F: −; β-NAG − P: +; I: +; C: −; Gf: −; α-F: −; β-NAG +

Plaque, dog Plaque, dog Plaque, bear, cat, coyote, dog, wolf, monkey, bovine Plaque, infected cat bite wounds, primates

P: +; I: +; C: +; Gf: most strains −; α-F: −; β-NAG different P: +; I: +; C: +; Gf: most strains −; α-F: −; β-NAG different P: +; I: −; C: −; Gf: weakly +; α-F: −; β-NAG +

P: +; I: +; C: −; Gf: −; α-F: −; β-NAG −

P: +; I: +; C: −; Gf: −; α-F: −; β-NAG +

P: +; I: −; C: −; Gf: weakly +; α-F: −; β-NAG + P: +; I: +; C: −; Gf: −; α-F: −; β-NAG −

P: +; I: +; C: +; Gf: most strains −; α-F: −; β-NAG different

P, pigment production; I, Indole; C, Catalase; Gf, Glucose fermentation; F, α-Fucosidase; β-NAG, N-acetyl-β-glucosaminidase

the culture is 7.5. Porphyromonas bacteria produce metabolic products such as acetic acid, propionic acid, isobutyric acid, and isovaleric acid, as well as traces of succinic acid. Originally, members of the Porphyromonas genus were described as asaccharolytic and further characterized by their production of porphyrins, heme-derived brown to black pigments, developing within 48 h of cultivation. Since Porphyromonas catoniae was included in the genus, the description was amended to include both nonpigmented and weakly saccharolytic species. P. bennonis, P. somerae, and P. uenonis are other species which are weakly pigmented only after prolonged cultivation on blood agar. Based on morphological, chemical, and biochemical criteria differentiation of the members of the genus was enabled by 16S rRNA gene sequencing and DNA–DNA relatedness studies.

Table 50.1 lists characteristic properties of all species of the Porphyromonas genus described so far.1,4,5,7–10,12,15,20,21,26,31 Epidemiology. Epidemiological data for Porphyromonas spp. are increasingly available. Porphyromonas spp. are considered to be part of flora of the normal oral and gastrointestinal tract as well as of the vagina.38,39 Porphyromonas spp. were found to be prevalent in abscesses, mostly of the head and neck; chronic otitis media, peritonitis, aspiration pneumonia, sinusitis, and periodontitis and were often isolated from mixed infections with other anaerobes and aerobic or facultative species.40 Porphyromonas and/or Prevotella spp. can be isolated from 20% to 30% of all anaerobic infections in children, and 10% of anaerobic bacteraemia.41–43 Studies on the composition of the subgingival microbiota in periodontitis patients have been performed in North American,

Porphyromonas

European, Japanese, and Latin American populations.14,44–50 Few studies are available addressing differences of the occurrence of P. gingivalis and P. endodontalis in different countries or different geographic regions.46–55 These studies suggest that there may be substantial differences in prevalence and levels of subgingival or endodontal species. Mean proportions of P. gingivalis ranged from 2% to 34% of the microbiota in Chilean, Colombian, Spanish, United States, Brazilian, and Swedish patients with chronic periodontitis.46,47 Using DNA–DNA checkerboard hybridization technique the detection frequency within Mexican subjects with chronic periodontitis reached 100% of all subjects with untreated chronic or aggressive periodontitis for P. gingivalis and 90% for P. endodontalis.45 P. endodontalis has been isolated from samples of primary endodontic infections or endodontal abscesses of Brazilian, American, Italian and, Korean provenance in comparative investigations.49,50,52–54 It has been concluded from these studies that differences of prevalence and proportion of P. gingivalis and P. endodontalis in different populations can occur. However, the small number of subjects as well as different subject-inclusion criteria may confound final statements.

50.1.2  Pathogenesis and Clinical Features This section describes the pathogenesis and clinical features for the seven Porphyromonas species that were isolated from human sources. All of them are associated with inflammatory processes according to the current state of knowledge. However, their virulence differs significantly on both the species and subspecies level. Porphyromonas have several virulence factors. Some of them enable the bacteria to establish in tissues by suppressing the immune defense of the host; others are responsible for provision of nutrients. The most important virulence factors are membrane-associated cysteine proteases (gingipains) together with further proteases. Hemagglutinins and hemolysins are responsible for the iron supply. The docking to host tissues is enabled by fimbria, which themselves carry important enzymes. A hydrophilic capsule provides protection. Capsular vesicles contain antigens, for example, heat shock proteins. Compared to other bacteria, the lipopolysaccharides (LPS) of Porphyromonas, especially of Porphyromonas gingivalis, have a low biological activity. Metabolic products, such as ammonia, organic acids, amines, and hydrogen sulfide are toxic for the host’s cells. Porphyromonas coaggregate with a large number of anaerobe and aerobe species. The Porphyromonas species most often isolated from human samples were P. asaccharolytica, P. endodontalis, and P. gingivalis. The pathogenicity of P. gingivalis has been investigated most extensively, and this species might be the most important pathogen of the genus. Little information is available on other species isolated from human sources, such as P. bennonis, P. catoniae, P. somerae, and P. uenonis. Porphyromonas is known to inhabit the oral cavity, colon, and small intestine as well as the urogenital system. Most often, however, they have been isolated from

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periodontal pockets and infected root canals. The share of anaerobic bacteria in periodontal pockets can reach up to 90%. Porphyromonas and Fusobacteria can dominate these habitats. Members of the genus have been found in numerous abscesses in various body regions. Porphyromonas spp. also appear in actinomycoses. There are indications that members of the Porphyromonas genus occur as commensal, opportunistic, as well as pathogenic bacteria. Infection can be passed horizontally, for example, between siblings and between parents, as well as vertically, from parents to children, and probably via smear infections and from anaerobe habitats of the environment.56 P. asaccharolytica.1 Pathogenesis: The physiology of P. asaccharolytica is very similar to P. endodontalis. The enzyme alpha-fucosidase distinguishes P. asaccharolytica from P. endodontalis. Together with P. gingivalis and P. endodontalis, this species is among those on which the original description of the genus Porphyromonas was based. Little is known about the virulence factors of P. asaccharolytica. The species is able to utilize human immunoglobulin G (IgG) as a substrate for growth. In contrast to P. gingivalis, only partial breakdown of IgG was observed.57 P. asaccharolytica shows activity of glycylprolyl protease, elastase-like activity, and phospholipase C activity. It has no trypsin-like activity and cannot hydrolyze type I collagen. The lectin WGA reacts strongly with the cell surface of P. asaccharolytica.6 OmpA proteins from P. asaccharolytica (OMP-PA), which are monomeric porins, have been found to induce release and expression of the proinflammatory cytokines IL-1-alpha, tumor necrosis factor (TNF) alpha, IFN-gamma, IL-6, and IL-10 in a murine model.58 P. asaccharolytica can contain plasmids59 and is susceptible to the nonoxidative killing mechanisms of human neutrophils.60 Clinical Features: P. asaccharolytica is a common medical pathogen. It has been isolated from numerous mucous membranes as well as from mostly intraabdominal abscesses with a rate of 41%.61 P. asaccharolytica is part of the healthy and infected vaginal flora, with an increased incidence in women with infections or other gynecological problems.62 This species is also associated with preterm deliveries63 and is often isolated from men with genital ulcers.64 It has also been shown to be associated with diversion colitis.65 Some case reports associate P. asaccharolytica with Lemierre’s disease.66 Fusobacterium spp. are known to cause Lemierre’s syndrome with Porphyromonas spp. as a possible accompanying species. P. asaccharolytica was identified as predominant bacterium in bone tissues, brain, liver, retroperitoneal, intraabdominal, chest wall, dental, and other abscesses in numerous further case reports.40,61,67 P. asaccharolytica has been described to trigger bacteremia via intravenous catheters.68 The pathogen has been isolated from patients with chronically inflamed maxillary sinuses67 in a mixed flora of aerobe and anaerobe species. P. asaccharolytica can contribute to diseases of the cardiovascular system, as in the case of an infected left atrial myxoma.69 It has also been isolated from the oral cavity. Its contribution to the pathogenesis of periodontitis is discussed controversially. According to

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Tran and coworkers, P. asaccharolytica does not seem to be part of the periodontopathic microbiota in humans.70 It was isolated from only 10% of patients suffering from juvenile periodontitis.71 In contrast, Rawlinson et al. found it to be the species most frequently identified in chronic periodontitis.72 Together with Fusobacterium nucleatum, Eubacterium lentum, and Peptostreptococcus micros, it was isolated from infected root canals.73 It is also frequently present in infected root dentin.74 P. asaccharolytica can be effectively eradicated from abscesses and mucous membranes by standard antibiotic therapy. Antibiotic resistances do not seem to be very common. Beta-lactamase activity has been observed only in some strains.61 In the treatment of periodontitis, root planning helps to reduce the frequency with which P. asaccharolytica can be isolated.75 P. bennonis.4 This is the most recently described species of the genus Porphyromonas. It was first observed in 2009 in samples harvested from abscesses in various parts of the human body where it cohabitated with anaerobes and aerobes. It was most frequently isolated in chronic infections of skin and soft tissues in the perirectal/buttock/groin regions. Fifteen percent of the strains produced β-lactamase. P. catoniae.5 The species was identified in samples of gingival crevices of patients with gingivitis, active periodontal site, juvenile periodontitis as well as from humans with healthy gums.76–78 Most frequently it is found in sites unaffected by destructive processes.34 It was detected in 70% of the infants aged 12 months, but only in 2% of 2-month-old babies, making it obviously an early colonizer of the oral cavity.14 P. endodontalis.1,6 Pathogenesis: P. endodontalis is a bacterium frequently found in the endodontium and in inflamed periapical tissues. As a strict anaerobe, it is adjusted to the conditions in the root canal and other anaerobe habitats. In order to survive in a community of organisms, P. endodontalis requires the porphyrin ring, preferably supplied as hemoglobin, as a growth supplement.79 The end products n-butyric acid, acetic acid, hydrogen sulfide, and methyl mercaptan are cytotoxic,80 for example, butyrate, a metabolic lipid byproduct causes G2/M phase arrests and inhibits protein synthesis.81 The bacterium is frequently encountered in mixed infections together with example, Fusobacterium nucleatum, P. gingivalis, Tannerella forsythia, Prevotella sp., Parvimonas micra (Peptostreptococcus micros), Selenomonas sputigena, or Campylobacter rectus.82,83 P. endodontalis binds hemoglobin that appears to serve as a sole carbon, iron, and nitrogen source.79,84 P. endodontalis produces proteases and peptidases. The proteases are able to degrade fibrinogen, fibronectin, and collagen type IV, and also cleave angiotensin. A proline aminopeptidase degrades bradykinin and vasopressin.85 The LPS from P. endodontalis was found to activate the complement system. P. endodontalis is able to degrade the complement factor C3.86,87 Degradation or inactivation of IgG is thought to play an important role in polymicrobial infections.57 LPS of P. endodontalis can rapidly induce the expression of the NF-κB-dependent gene interleukin-8 (IL-8) in dental-pulp stem cells (DPSCs), pulp fibroblasts, and osteoblasts.88,89 It can upregulate the receptor activator of NF-κB ligand

Molecular Detection of Human Bacterial Pathogens

(RANKL) production in osteoblasts and is thus involved in developing apical periodontitis through the stimulation of RANKL production and activation of osteoclasts.90 When human dental-pulp (HDP) cells were stimulated by LPS from P. endodontalis, the production of IL-6 always preceded that of IL-1beta. Since the IL-6 production was observed even in the presence of an IL-1beta receptor antagonist, it was concluded that IL-6 production was independent of the IL-1beta molecule in LPS-stimulated HDP cells.91 LPS released from the infected root canal triggers the synthesis of IL-1 alpha and TNF-alpha from macrophages and is capable of stimulating PMNs.92 These proinflammatory cytokines upregulate the production of MMP-1 by macrophages to promote periapical bone resorption.93 P. endodontalis also was found to elevate MMP-2 production in both human pulp and PDL cell cultures.94 Via IL-8, a potent inducer of neutrophil chemotaxis and activation, which also has a proangiogenic effect, P. endodontalis stimulated the vascular network coincident to progression of the inflammation through the stimulation of vascular endothelial growth factor (VEGF).95 In addition, proinflammatory cytokines and black-pigmented Bacteroides may be involved in developing pulpal inflammation through the stimulation of IL-6 production.89 P. endodontalis also was found to be able to induce the antiinflammatory cytokine IL-10 but not TNF-alpha, IL-12, or IFN-gamma.96 The activation of COX-2 may contribute to host degradative pathways in the pathogenesis of microbial-induced pulpal/periapical inflammation.97 In a rat model, the early stage of pulpal inflammation induced by P. endodontalis was accompanied by a shift of the CD4+:CD8+ ratio toward CD8+. The increase in IL-2 and IFN-gamma suggests a Th1 reaction.55 P. endodontalis enhances tissue plasminogen activator (t-PA) production in human pulp, osteoblasts, and gingival cells.98,99 Human coronary artery endothelial cells (HCAEC) and coronary artery smooth muscle cells (CASMC) can be invaded by P. endodontalis.100 Clinical Features: P. endodontalis has been isolated frequently in high counts from primarily and secondarily infected dental root canals, chronic periapical lesion, and submucous abscesses of endodontal origin.101–105 The prevalence of the pathogen ranges between 10% and 70%,106,107 depending on the methodology applied. Investigations comparing molecular detection methods with cultivation have shown higher prevalences for the former.108 P. endodontalis can be present in symptomatic and asymptomatic cases, with the prevalence being higher in acute apical inflammations.101,108 High levels of P. endodontalis were detected in carious dentine and in pulpal inflammation stages, from minimally inflammatory change to inflammatory degenerative change.109 It is occasionally found on healthy oral mucosa and, more frequently, in gingivitis.110 The species has been associated with periodontitis.111 Differences in virulence between strains have been related to capsular material. On the basis of different types of capsules, three serotypes have been described.112

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Porphyromonas

P. endodontalis is sensitive to a wide range of antibiotics, including penicillin, tetracycline, and metronidazole.112 P. gingivalis.1 Pathogenesis: P. gingivalis is the most frequently found species of the genus and is the one most comprehensively investigated. This bacterium has its habitat in the human oral cavity and is associated with initiation and progression of a group of chronic inflammatory diseases of the gingival and supporting periodontal tissues. P. gingivalis possesses a wide range of virulence factors, helping it to settle in the oral ecosystem. Fimbriae are effective for adhesion on tissues, bacterial coaggregation,113–117 and for initiating hemagglutination reactions.118,119 The hemoglobin released by destruction of erythrocytes by hemolysins is made accessible to the nutritional supply.120,121 Cell-surface anionic polysaccharides provide resistance to complement-mediated killing.122 The LPS of P. gingivalis may contribute to inhibition of the innate immune response.123,124 Different proteinases, particularly cysteine proteinases (gingipains), are considered the prominent virulence factor of the pathogen. These proteinases are responsible for the high proteolytic activity of the bacterium. P. gingivalis can survive inside as well as outside epithelial or endothelial cells.125–127 It seems to be able to undermine immune clearance via exploitation of the complement receptor-/CR3 in monocytes/macrophages.128 In biofilms, coaggregations of P. gingivalis, T. forsythensis, and T. denticola, or of P. gingivalis and A. actinomycetemcomitans, or F. nucleatum, are positively correlated with a progressive course of disease.126,129,130 The potential key virulence factors of P. gingivalis are briefly listed in the following (Figure 50.1): (i) Fimbriae: Fimbriae mediate bacterial adherence to and entry into periodontal cells and contribute to coaggregation with species within the oral ecosystem.131,132 P. gingivalis produces fimbriae of between 0.5 and 1.6 µm in length and approximately 5 nm in width.133,134 Most strains of P. gingivalis

V F

C

FIGURE 50.1  Transmission electron image of Porphyromonas gingivalis stained with ruthenium red. V, extracellular vesicles; C, capsule; F, fimbriae. Bar = 330 nm.

produce fimbriae consisting of fimbrillin subunits of 41–49 kDa.133 The gene encoding the P. gingivalis fimbriae (fimA) is resident on the chromosome as a single copy. All strains so far examined contain this gene.135,136 Nucleic acid sequencing of fimA genes revealed five distinct clusters among P. gingivalis strains.137 Fimbriae have an antigenic effect; proteases (gingipains) placed on them release cryptic ligands and promote hemagglutination.138 They mediate the binding to a number of molecules and structures, such as saliva molecules, epithelial cells, other bacteria, fibrinogen, fibronectin, and lactoferrin.139–144 In addition to adhesion, mediation fimbriae are supposed to possess chemotactic properties and to discharge cytokines. Intact fimbriae, fimbrillin-specific peptides, or synthetic fimbrial peptides can stimulate the production of fibroblast-derived thymocyte-activating factor from human gingival fibroblasts, interleukin (IL)-1, IL-6, IL-8, neutrophil chemotactic factor KC, and tumor necrosis factor (TNF)-alpha from mouse peritoneal macrophages or in human peripheral blood monocytes.136 (ii) Minor fimbriae: A second type of fimbriae was isolated and characterized, which was found to have a higher molecular weight of the fimbrillin, 67 kDa. These fimbriae seem to play an important role in host bone loss.145 (iii) Membrane-associated proteinases: Thiol-activated trypsin-like cysteine proteinases (gingipains) have been recognized as major proteolytic enzymes of this organism. RgpA and RgpB (gingipain R: encoding genes: rgpA and rgpB) are arginine-specific proteinases, Kgp (gingipain K, kgp gene) is lysine-specific. RgpA and Kgp contain separate catalytic and adhesion/hemagglutinin domains, while RgpB has only a catalytic domain. The most important tasks of RgpA and RgpB are activation of the kallikrein-kinin system, inducing enhanced vascular permeability, and activation of the blood coagulation system. The activation of the kinin system is involved in breaking the vascular barrier that permits dissemination of P. gingivalis.146 Kgp is the most potent fibrinogen/ fibrin degrading enzyme and degrades also hemoglobin to heme. RgpA activates coagulation factors and degrades fibrinogen/fibrin. Gingipains degrade macrophage CD14, thus inhibiting activation of leukocytes through the lipopolysaccharide (LPS) receptor.147 The proteinases degrade complement factor C3, preventing the deposition of C3b on the bacterial cell surface. The degradation of C5 leads to generation of a chemotactic C5a-like fragment.148 This contributes to significant leukocyte infiltration while simultaneously inactivating their C5a and FMLP receptors. This may lead to P. gingivalis being protected from phagocytosis by PMN.149 C5a receptor cross-talks with TLR signaling pathways and inhibits IL-12.

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(iv)

(v)

(vi)

(vii)

Molecular Detection of Human Bacterial Pathogens

This led to the assumption that P. gingivalis uses different complement-related mechanisms for escaping IL-12–mediated clearance.88,128,150 Gingipains also have collagenolytic activity and degrade or inactivate inflammatory cytokines IL-6, IL-8, TNF-alpha, and IFN-gamma. The activation of host derived collagenases causes further tissue damage. Two other proteinases (Trp, PrtT) have MMP-activating or hemagglutinating properties. Dipeptidyl aminopeptidase IV (DPPIV) may act as a virulence factor by contributing to the degradation of connective tissue.151 Hemagglutinins and hemolysins: P. gingivalis has essential requirements for both iron and protoporphyrin IX, which it preferentially obtains from heme. Several distinct hemagglutinins (HagA– HagC and further) have been identified.137,152,153 The proteinase Kgp specifically binds and cleaves hemoglobin.154 Outer membrane receptors (HmuR, HmuY) are responsible for heme utilization.155–157 Lipopolysaccharide: P. gingivalis LPS differs from the LPS of other bacteria. It induces in vivo endothelial cell expression of E-selectin at significantly lower levels compared to enterobacterial LPS, and inhibits E-selectin upregulation by other bacteria in vitro.123,158 This may result in diminished neutrophil adhesion to endothelial cells. P. gingivalis expresses heterogeneous LPS, including atypical lipid A structures, that trigger TLR2 signaling, and weakly stimulate or antagonize TLR4 activation.159,160 This may suppress TLR-mediated immunity.159 P. gingivalis LPS upregulates the IL-1R–associated kinase (IRAK)-M and relatively poorly induce IL-1beta and TNF-alpha compared with E. coli LPS.125,161,162 The binding of P. gingivalis LPS to LPS binding protein (LBP) is weak compared to E. coli LPS.163 In mice, the LPS induces apoptosis before a specific immune response to eliminate early nonspecific activated lymphocytes. This suggests a direction of the periodontal inflammation infiltrate toward a Th2 reaction.164 Capsule: Most P. gingivalis strains are able to produce capsular polysaccharides. This results in reduced binding of P. gingivalis to PMN and inhibited phagocytosis.165,166 Six capsular serotypes of P. gingivalis have been described, and a number of nontypeable capsule serotypes appear to exist. In a mouse model, the capsular serotypes were more virulent than nonencapsulated strains.167–169 Outer membrane vesicles (MV): P. gingivalis secretes outer membrane vesicles that contain major virulence factors, including the proteases Arggingipain (Rgp) and Lys-gingipain (Kgp). In vitro, MVs enter human epithelial cells via a lipid raftdependent endocytic pathway and survive within the endosomes for an extended period. MV-associated

gingipains degrade cellular functional molecules such as TfR and paxillin/FAK, resulting in cellular damage, indicating that P. gingivalis MVs are potent vehicles for the transmission of virulence factors into host cells and are involved in the etiology of periodontitis.170,171 (viii) Toxic metabolites: Organic acids, amines, ammonia, hydrogen sulfide, or butyric acid are inducers for apoptosis in B cells and T cells.172,173 Clinical Features: P. gingivalis has been strongly associated with the pathology of chronic periodontitis.104 Periodontitis is a multibacterial infection that affects the tooth-supporting tissues by destroying connective tissues and alveolar bone. A cyclical course is characteristic for this disease. The presence of specific bacteria at high levels in the subgingival plaque as well as the individual susceptibility to periodontal disease influence the progression of periodontitis. Important influence factors are cigarette smoking and poorly controlled diabetes mellitus. Genetic determinants and other common diseases are discussed to be modifiers for the onset and severity of periodontitis.174 The human oral cavity is composed of multiple epithelial and mucosal surfaces, as well as the calcified hard-substance enamel and the hard-tissue dentin. All oral surfaces are constantly moistened by saliva. Periodontopathogenic bacteria use saliva as a transmission medium. Various studies have shown that an organism is colonized by only one or at least one dominating P. gingivalis genotype. A large number of different strains could be isolated from different patients, supporting the hypothesis that P. gingivalis is to be considered an opportunistic pathogen, whose pathogenicity, however, cannot be assigned to a particular genotype. Precondition for the survival of the bacterium is its adhesion to epithelia or hard surfaces. The prevalence of P. gingivalis is lower in persons with healthy periodonts than in patients with periodontal diseases.175–181 P. gingivalis can be isolated from periodontal pockets, supragingival plaque, root canals, saliva, tongue, and the buccal mucosa, but also from the tonsils.161,180,182–184 Most frequently, the bacterium is found in deep periodontal pockets, where it also yields the largest counts. It has also been isolated from abscesses of the periodontium and endodontium. Its DNA was found in artheromatous plaques together with the DNA of other periodontal pathogens,185 corneal scrapings,186 in positive transient bacteremia subjects,187 cavernous sinus thrombophlebitis,185 from a severe brain abscess,188 and a tubal-ovarian abscess.189 P. gingivalis has been found in reduced prevalence in edentulous elderly.176 There is increasing evidence from epidemiological studies, case studies, and scientific observations that periodontal disease represents a significant risk factor for cardiovascular and cerebrovascular disease. Potentially periodontal diseases may be important for preterm delivery as well as low birth weight of newborns. Recent studies suggest that there is a bidirectional relationship between diabetes mellitus and periodontal disease.190 Periodontal disease can compromise blood sugar regulation in diabetes on the one

Porphyromonas

hand. On the other hand, diabetes has been identified as a significant risk factor for periodontal disease.191 The occurrence of P. gingivalis has been found to be associated with stroke and arteriosclerosis, myocardial infarction, coronary heart disease, preterm births, diabetes mellitus, and respiratory diseases.125,128,192,193 P. somerae (P. levii-Like Organisms [PLLO]).7 P.  somerae is associated with chronic skin and soft-tissue or bone infections, especially in the lower extremity. PLLO, now designated as P. somerae, have been isolated from diverse human clinical samples,194 for example, from the vagina, from patients with chronic otitis media, and one case of bacterial vaginosis. A systemic predisposition such as diabetes mellitus, neuropathy or peripheral vascular disease seems to facilitate infections in which P. somerae is the predominant bacterium.7 The species seems always to cohabitate with other anaerobes, such as anaerobic cocci, Bacteroides fragilis group, or Prevotella spp. About 20% of the strains are β-lactamase producers, and some strains have been found to be resistant to clindamycin. P. uenonis (P. endodontalis-Like Organism [PELO]). 8,9 P. uenonis appears most often in samples of intestinal origin, suggesting the human gut to be its preferred habitat. When isolated from infectious material, it was always accompanied by other anaerobes or aerobes. Some strains have been found to be resistant to penicillin G, ampicillin, and trimethoprim. P. uenonis seems to have a low virulence as it is exclusively found in mixed culture but not in blood cultures. It is not associated with serious infections.

50.1.3  Diagnostics 50.1.3.1  Conventional Techniques Conventional techniques for the detection of Porphyromonas often have a lower sensitivity compared with molecular methods. Primarily, this relates to the limited viability of Porphyromonas outside anaerobic habitats. The transport of the samples from patient to a microbiological laboratory is therefore a particular challenge.195,196 Once the bacteria have been successfully isolated, antimicrobial resistances can be determined on the basis of cultivation.197 Cultivation Techniques and Biochemical Identification. Cultivation still plays an important role in the detection of Porphyromonas. Samples are mostly mixed populations of aerobic and anaerobic microorganisms. Cultivation is performed by an anaerobic technique on blood agar198; identification, by the detection of specific enzyme patterns.199 If bacteria have been successfully isolated, further differentiations on the species and subspecies level can be performed on the basis of cultivation using different methods of characterization. Immunofluorescence. Specific poly- or monoclonal antibodies can be used for the direct detection and quantification of P. gingivalis in plaque samples as well as for its intracellular detection in gingival epithelial or endothelial cells.200–203 Fluorescent dyes coupled to the antibodies make the targeted bacteria directly visible.

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Matrix-Assisted Laser Desorption/Ionization Timeof-Flight Mass Spectrometry (MALDI-TOF-MS). MALDI-TOF-MS is a soft ionization method, which allows the phenotypic identification of different cultured bacteria.204–206 Ions are separated and detected according to their masses and charges. Each mass peak corresponds to a molecular fragment released from the cell during laser desorption. Bacteria, including Porphyromonas, can be identified by comparing their mass spectrum with those obtained from known reference strains using methods of pattern analysis.198,205,207 Other Methods. The BANA chair-side test was a diagnostic method for the detection of bacterial enzymes. Trypsinlike proteinases, produced among others by P. gingivalis were detected by blue coloration of a test stripe. However, the test had an insufficient specificity.208 Serological diagnostics detected antibody reactivity in the serum or sulcus fluid against specific bacterial antigens.209 Restriction endonuclease analysis (REA) and ribotyping were successfully applied for the typing of P. gingivalis.210 Analysis of electrophoretic mobilities of 16 metabolic enzymes of 100 P. gingivalis strains from humans and animals revealed 78 different multilocus enzyme electrophoresis (MLEE) types.182 50.1.3.2  Molecular Techniques Nucleic acid–based methods are indispensable for the detection of Porphyromonas, particularly of P. gingivalis, as their introduction has enabled a fast and inexpensive diagnostic. They are clinically used in cases of severe or aggressive periodontitis for diagnosis, reevaluation after a treatment phase, and during recall. The most important molecular methods for the detection and quantification of P. gingivalis include hybridization using species-specific DNA probes, conventional end-point PCR as well as quantitative end-point PCR, or quantitative real-time PCR using DNA intercalating dyes or fluorogenic DNA probes. The majority of current methods for detection and enumeration of P. gingivalis are based on 16S ribosomal RNA operon sequences.211–213 Usage of the rRNA operon offers the advantage of every bacterial cell harboring several template copies. The P. gingivalis strain W83 is known to have 4 rRNA operons (The Ribosomal RNA Operon Copy Number Database, http://ribosome.mmg.msu.edu/rrndb/index.php). Single-copy gene assays based on species-specific virulence factor genes have been developed for P. gingivalis using sequences of the Arg-gingipain,214 the waaA (3-deoxy-d-manno-oct-2-ulosonic acid transferase) gene, or the fimA genes.215 PCR Techniques. PCR is an established method for the detection of P. gingivalis.216,217 Once its specificity has been tested, it has low demands on sampling and sample transportation, since neither living nor cultivable bacteria is required. Bacterial species and subspecies can be detected or quantified by means of different PCR techniques. Quantitative PCR also enables quantification of the entire number of copies of bacterial DNA contained in a sample.218 Ribosomal and single-copy genes can be used as target sequences. The genes of 16S rRNA, OMP, FimA, rgpB, or PrtC are frequently used for P. gingivalis.216,219 Classic nonquantitative PCR, having

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a detection level of less than 100 cells, is often used in studies on the prevalence of this bacterium.220 Another technique used was multiplex PCR using three 16S rRNA speciesspecific forward primers and a conserved reverse primer for the simultaneous detection of A. actinomycetemcomitans, T. forsythensis, and P. gingivalis.221 While no quantitative information is obtained by means of classic PCR techniques, the number of bacterial cells can be determined by means of quantitative PCR. Competitive and real-time PCR techniques were developed for Porphyromonas.213,215,219 In competitive PCR, inhibitors of amplification are easily recognized by preventing the formation of the competitor product. In contrast, in classic as well as in real-time PCRs, inhibitors of Taq polymerase and the presence of other bacterial or human DNA may lead to false-negative results. Real-time PCR techniques, which are routinely used for quantification of Porphyromonas, offer a fast quantification and monitoring of the amplification through DNA intercalating dyes (SYBR green), as well as control of the specificity by dye-labeled specific probes. The comparison of PCR with subsequent reverse hybridization and checkerboard DNA–DNA hybridization for the identification of bacterial species in subgingival plaque samples showed that both tests could distinguish samples from healthy and periodontitis subjects.222 Different PCR based tests for detection and quantification of P. gingivalis are commercially available.222,223 The clonal diversity of the taxon P. gingivalis was studied using molecular fingerprint techniques, like arbitrarily primed (AP) PCR, repetitive extragenic palindromic (REP),210 and random amplified polymorphic DNA (RAPD)-PCR.224 RNA Hybridization. Identification and quantification of bacteria can be performed by direct hybridization of specific probes with the ribosomal RNA. Since many 16S rRNA copies exist per cell, no amplification of the target nucleic acids is necessary. However, since bacterial RNA is rapidly degraded after cell death, only living bacteria are detected. Hybridization of RNA requires a buffered transport system that protects bacteria from death but, at the same time, inhibits bacterial growth. The best-known hybridization test currently available on the market is IAI PadoTest 4.5.225,226 DNA Hybridization. The “checkerboard” DNA–DNA technique uses DNA probes on a single support membrane for hybridizing large numbers of DNA samples. Denatured DNA from up to 43 samples can be fixed in separate lanes on a single membrane mounted in a Miniblotter 45. The membrane is then rotated by 90° in the same device, which enables simultaneous hybridization with 43 different DNA probes. Hybridizations are performed with either digoxigenin­labeled whole genomic probes or 16S rRNA-based oligonucleotide probes directly conjugated to alkaline phosphatase. The method permits the simultaneous determination of the presence of multiple bacterial species in single or multiple dental plaque samples, thus suggesting its usefulness for a range of clinical or environmental samples.227 A modification of the method, the reverse-capture checkerboard hybridization, uses RT–PCR as a first step to amplify

Molecular Detection of Human Bacterial Pathogens

16S rRNA genes. After that, hybridization with oligonucleotide probes allows to differentiate between closely related species or even subspecies.228 Other Molecular Techniques. PCR is often the basic technology for more complex applications. For example, there is a multilocus sequence typing (MLST) scheme for P. gingivalis. By a combination of PCR and DNA-sequencing, the heterogeneity of a set of target genes is used for a strain differentiation on the subspecies level. The following target genes are used for P. gingivalis: gdpxJ, pga, hagB, mcmA, recA, ftsQ, and pepO.229 PCR is also the basic technology for the human oral microbe identification microarray (HOMIM) developed by The Forsyth Institute in Boston. HOMIM represents a high throughput technology to examine complex oral microbial diversity in a single hybridization. About 300 oral bacterial species (http://mim.forsyth.org/bacteria.html), including the “uncultivables” and Porphyromonas spp. are simultaneously detected.230 A DNA-chip has been developed231 allowing a semiquantitative detection of periodontal bacteria, including P. gingivalis. Bacterial DNA has to be extracted, and part of the 16S rRNA gene is amplified by PCR using highly conserved primers labeled with a fluorophore (Cy5). Without further treatment, samples are hybridized to the DNA-chip, which is spotted with strain-specific DNA probes. Terminal restriction fragment length polymorphism (T-RFLP) and denaturating gradient gel electrophoresis (DGGE) are alternative molecular approaches that allow the assessment of diversity of oral microbiota and rapid comparison of community structures. These techniques can contribute to the identification of yet unknown Porphyromonas spp.232,233

50.2  METHODS 50.2.1  Sample Preparation Sample taking and preparation is not critical for cultured Porphyromonas. Clinical samples of Porphyromonas, mostly of oral origin, are very complex and may contain several hundred other bacteria species. In addition, oral samples are often contaminated by saliva, pus, and above all, blood. As hemoglobin strongly inhibits PCR and other enzyme-catalyzed reactions, an efficient DNA preparation plays a decisive role. Sampling Methods.184,234–237 The sampling strategy depends on the aim of the investigation and the method of analysis applied. In DNA based molecular techniques, cotton swabs or paper points can be shortly stored in 1.5 mL tubes at 4°C or at −20°C for longer periods of time. Aspirates from abscesses, plaque samples or whole saliva should be stored frozen. Large samples have to be centrifuged, and the pellets have to be resuspended in suitable smaller volumes, for example, Tris-EDTA buffer, before further processing. Smaller sample volumes can be suspended directly in 100–200 µL Tris-EDTA buffer. Sampling techniques routinely used for Porphyromonas are described in the following: Cultures: One colony is to be scraped from agar plates and suspended in 200 µL Tris-EDTA buffer.

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Porphyromonas

Mucosal (Skin, Abscess) Swabs: Streak the surface of the mucosa vigorously back and forth with a sterile swab. Resuspend the sample in 1 mL Tris-EDTA buffer. Saliva: Salivary flow is stimulated by paraffin chewing for 2 min. Whole saliva samples are collected by expectoration. Saliva samples are centrifuged at 10,000 × g for 5 min, the supernatant has to be discarded, and the samples are resolved in 200 µL Tris-EDTA buffer. Oral Lavage: Rinse with 10 mL sterile saline for 30 s and collect samples by expectoration. Then follow the procedure described for saliva. Subgingival Plaque (Paper Points; Figure 50.2): Isolate sample sites with cotton rolls. Supragingival plaque has to be removed, and a sterile paper point (iso 40) is inserted into the pocket until meeting with resistance. After 10 s, the point is removed and stored in a 1.5 mL tube at −20°C until further investigation. There are site-specific and pooled-sample strategies. Pooled samples from the (minimally three) deepest active periodontal pockets provide representative information about the subgingival microflora of the whole oral cavity which is relevant for adjunctive systemic antibiotic therapy.184 Subgingival Plaque (Curette): Sample sites are isolated with cotton rolls. Supragingival plaque has to be removed and discarded. A Gracey curette is inserted to the base of the pocket or sulcus, and as much plaque as possible is removed with a single stroke. The curette plus sample has to be shaken in a 1.5 mL tube containing 200 µL Tris-EDTA until no visible plaque remains on the curette. Supragingival Plaque (Curette): Sample sites are isolated with cotton rolls. A Gracey curette stroke is performed on the surface of the tooth without inserting the curette into the periodontal pocket. DNA Preparation. The method of DNA preparation depends on sample characteristics and the detection method used. Inflammation exudate, blood, saliva, or human DNA in the sample can also influence the results. Many different

FIGURE 50.2  Sampling of subgingival plaque with a paper point at the mesio-buccal aspect of a right maxillary central incisor.

DNA preparation protocols have been described that are subject to constant modification and improvement. There are two basic sample preparation strategies: Simple preparation for cultured bacteria: Suspend a single bacterial colony in 50 µL of 0.7% (w/v) sodium chloride solution, and dilute this suspension subsequently 1:100 again in 0.7% sodium chloride; use 5 µL of this dilution as template for PCR.229 Complex preparation of clinical samples: Different kits for DNA preparation are available and commonly used in clinical and laboratory studies. A detailed DNA-isolation protocol is recommended including sophisticated lysis by lysozyme and Proteinase K and DNA precipitation by isopropanol/ethanol by The Forsyth Institute: (http://mim.forsyth. org/samplepreparation.html). For quantitative competitive PCR of bacteria in plaque or saliva, the following protocol is used238,239: Add 50 µL of glass beads with diameters of 100–250 µm to 200 µL aliquots of the sample and vortex for 4 × 30 s at 30 Hz. Add 100 µL of a buffer containing 0.01 M Tris, 0.005 EDTA and 0.5% SDS and 1 µL lysozyme (50 mg/mL) to 15-µL aliquots of the supernatant. Add appropriate amounts of DNA competitor. Incubate the mix for 15 min at 4°C. Add 5 µL of proteinase K (19 mg/ mL), and then incubate for 1 h at 50°C. For DNA precipitation, add 10 µL of sodium acetate and 500 µL of ethanol (96%). Incubate at −80°C for 30 min then centrifuge for 30 min at 4°C and 10,000 g. Next, 500 µL of 75% ethanol has to be followed by centrifugation for 15 min at 4°C and 10,000 × g. Redissolve the pellet in 15 µL of nanopure water.

50.2.2  Detection Procedures 50.2.2.1  Species-Specific PCR213 Primer Sequences (5′–3′): Target gene: 16S rRNA gene, forward (pg1): 5′-GGG ATT GAA ATG TAG ATG ACT GAT G-3′, reverse (pg2): 5′-CCT TCA GGT ACC CCC GAC T-3′, 488 bp. Performed in a 50 µL reaction volume in 0.2 mL tubes, 2 µL of sample (template) supplemented with 40 pmol of each primer, 0.1 µmol MgCl2, 2.5 nmol each of deoxynucleoside triphosphate, 1.25 U Thermus aquaticus polymerase, 5 µL 10× Taq buffer, and water to 50 µL. PCR amplification, performed in a standard DNA thermal cycler, initial denaturation step at 95°C for 10 min followed by 40 cycles of denaturation at 95°C, primer annealing at 55°C and extension at 72°C, for 1 min each, and a final step at 72°C for 10 min. The amplified material was stored at −20°C. 50.2.2.2  Quantitative Real-Time PCR Primer Sequences (5′–3′): See 50.2.2.1. LightCycler assay with kinetic analysis (fluorescence threshold method) (238, 239): PCR is performed with 20 µL reaction mixture containing 2 µL LightCycler-DNA Master SYBR Green I (Roche Molecular Biochemicals), 6 mM MgCl2, 2–4 µL DNA template, and 0.5 mM of both forward and reverse primers. For amplification, 45 cycles of 94°C for 0 s, 55°C for 10 s, and 72°C for 20 s (temperature transition

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of 20°C/s), preceded by denaturation at 94°C for 3 min, are used, and the fluorescence reading is taken at 72°C. The melting curve analysis is performed with continuous fluorescence reading between 65°C and 95°C with a transition rate of 0.1°C/s. For the kinetic analysis, the second derivative maximum method is used. 50.2.2.3  Q  uantitative Competitive PCR (QC–PCR)215,218,240 Primer Sequences (5′–3′): target gene: 16S rRNA gene P. gingivalis, 450 bp PCR product, primer see Section 50.2.2.1 and pg-hy: 5′-GGGATTGAAATGTAGATGACTGATG TCAGCTCGTGCCGTGAG-3′. This technique is an end-point PCR. Quantification of bacteria by PCR is carried out by coamplification of a homologous competitor. In order to facilitate the separation of the PCR products obtained from the sample and the homologous competitor, the latter is designed shorter than the sample template. Quantitative competitive PCR can be performed in a conventional block thermal cycler without using intercalating DNA dyes. This technique has been described for P. gingivalis and other bacteria and uses the genes of the 16S rRNA218,240,241 or the fimA gene.215 The sample (template) (2–5 µL) is supplemented with aliquots of competitors corresponding to 10, 100, or 1000 bacteria, respectively; 40 pmol of each primer; 0.1 µmol MgCl2; 2.5  nmol of each deoxynucleoside triphosphate; 1.25 U Thermus aquaticus (Taq) polymerase; 5 µL 10× Taq buffer; and water to 50 µL. PCR amplification, performed in a standard DNA thermal cycler, included an initial denaturation step at 95°C for 10 min, followed by 40 cycles of denaturation at 95°C; primer annealing at 55°C and extension at 72°C, for 1 min each; and a final step at 72°C for 10 min. The amplified material is stored at −20°C. Amplicons are detected by electrophoresis of 10 µL of PCR product on a 2% agarose gel. After staining with ethidium bromide DNA is quantified videodensitometrically. Counts of bacteria are calculated considering the amounts of 16S rRNA gene copies for the bacterial species investigated. Competitive template construction: The competitor (407 bp) is obtained by PCR using the reverse primer (pg2) together with a hybrid primer (hy-pg) that binds with its 3′ part inside the P. gingivalis 16S rRNA and contains the sequence of the reverse primer at its 5′-terminus (Figure 50.3). Alternatively, competitive templates can be constructed by cloning appropriate DNA fragments into plasmids constructing a heterologous P. gingivalis competitive template.215

50.3  CONCLUSIONS AND PERSPECTIVES The fast development of molecular methods has brought about a specification of the taxonomic position of the genus Porphyromonas. Molecular approaches independent of cultivation have revealed the diversity, for example, of human oral microbiota and the existence of a large number of asyet-to-be-cultured Porphyromonas spp. on the basis of 16S rRNA genes.242 In addition, molecular methods enable the

Molecular Detection of Human Bacterial Pathogens 16S rRNA gene from P. gingivalis pg1 hybrid primer (hy-pg)

pg2 PCR

homologous competitor (407 bp)

FIGURE 50.3  Principle of synthesis of the homologous competitor. With P. gingivalis 16S rRNA as template, the competitor DNA was obtained by PCR using the forward primer (pg1) together with the hybrid primer (hy-pg) that binds with its 3′-part inside the template and contains the sequence of the reverse primer (pg2) at its 5′-terminus.

culture-independent detection of Porphyromonas spp. on the species as well as on the subspecies level and their association with a number of diseases. The species P. asaccharolytica, P. endodontalis, and especially, P. gingivalis have been the most thoroughly investigated so far. Surely, further species will be detected and classified as Porphyromonas. Molecular techniques have also given important insights into the interactions between bacterium and host of the genus Porphyromonas, particularly P. gingivalis. Numerous virulence factors are known whose functions in vitro have been largely elucidated. P. gingivalis may be a key species in the oral microbial community. It may be able to modulate innate host defense for surviving in tissues intra- as well as extracellularly.243 The behavior of this organism in periodontal pockets may be unobtrusive or more aggressive. It still has to be finally clarified whether individual strains embody both characters or if different subspecies exist that are particularly virulent.125 A random distribution of potential virulence factors of P. gingivalis detected in strains of different geographical origin led to the conclusion, that P. gingivalis may have a nonclonal population structure characterized by frequent recombination.244 However, it is becoming increasingly clear that specific host factors play a decisive role in the clinical outcome. Hybridization techniques and PCR methods are widely used in clinical science and practice. However, the use of molecular methods for diagnosis and therapy planning is still limited. The different molecular techniques have different limits of detection. PCR, in particular nested PCR, has a very low detection limit. Combination of PCR and hybridization techniques can detect 102–104 bacteria. The measured prevalence of Porphyromonas may differ according to the applied method. This makes the correlation of molecular detection results with clinical diagnoses complicated. Above all, the amount of P. gingivalis present in the periodontal pocket seems to be decisive for the progression of oral periodontal diseases,245 which may also be true for other species of the genus Porphyromonas. For clinical application, the detection of P. gingivalis is a useful tool for motivating the patient by visualizing the infection. As P. gingivalis cannot

Porphyromonas

be completely eradicated by mechanical periodontal therapy, the detection of this pathogen in high counts or percentages of the periodontal flora can support the decision to use systemic antibiotics adjunctively.246 The application of quantitative molecular techniques offers advantages in comparison to nonquantitative detection. No molecular technique can yet predict, however, the pathogenic potential of an indigenous P. gingivalis population or a bacterial community for its host.

REFERENCES











1. 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. 2. 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. (Editors), Bergey’s Manual of Systematic Bacteriology, Vol. I, pp. 604–631, The Williams & Wilkins Co., Baltimore, MD, 1984. 3. Finegold, S.M., and Barnes, E.M., Report of the ICSB taxonomic subcommittee on Gram-negative anaerobic rods, Int. J. Syst. Bacteriol., 27, 388, 1977. 4. Summanen, P.H., Lawson, P.A., and Finegold, S.M., Porphyromonas bennonis sp. nov., isolated from human clinical specimens, Int. J. Syst. Evol. Microbiol., 59, 1727, 2009. 5. Willems, A., and Collins, M.D., Reclassification of Oribaculum catoniae (Moore and Moore 1994) as Porphyromonas catoniae comb. nov. and emendation of the genus Porphyromonas, Int. J. Syst. Bacteriol., 45, 578, 1995. 6. Van Steenbergen, T.J.M. et al., Bacteroides endodontalis sp. nov., an asaccharolytic black-pigmented Bacteroides species from infected dental root canals, Int. J. Syst. Bacteriol., 34, 118, 1984. 7. Summanen, P.H. et al., Porphyromonas somerae sp. nov., a pathogen isolated from humans and distinct from Porphyromonas levii, J. Clin. Microbiol., 43, 4455, 2005. 8. Conrads, G. et al., 16S-23S rRNA gene internal transcribed spacer sequences for analysis of the phylogenetic relationships among species of the genus Porphyromonas, Int. J. Syst. Evol. Microbiol., 55, 607, 2005. 9. Finegold, S.M. et al., Porphyromonas uenonis sp. nov., a pathogen for humans distinct from P. asaccharolytica and P. endodontalis, J. Clin. Microbiol., 42, 5298, 2004. 10. Collins, M.D. et al., Phylogenetic analysis of members of the genus Porphyromonas and description of Porphyromonas cangingivalis sp. nov. and Porphyromonas cansulci sp. nov., Int. J. Syst. Bacteriol., 44, 674, 1994. 11. Allaker, R.P. et al., Dental plaque flora of the dog with reference to fastidious and anaerobic bacteria associated with bites, J. Vet. Dent., 14, 127, 1997. 12. Love, D.N. et al., Porphyromonas canoris sp. nov., an asaccharolytic, black-pigmented species from the gingival sulcus of dogs, Int. J. Syst. Bacteriol., 44, 204, 1994. 13. Su, Y. et al., Changes in abundance of Lactobacillus spp. and Streptococcus suis in the stomach, jejunum and ileum of piglets after weaning, FEMS Microbiol. Ecol., 66, 546, 2008. 14. Kononen, E. et al., Establishment of oral anaerobes during the first year of life, J. Dent. Res., 78, 1634, 1999. 15. Love, D.N. et al., Description of Porphyromonas circumdentaria sp. nov. and reassignment of Bacteroides salivosus (Love, Johnson, Jones, and Calverley 1987) as Porphyromonas (Shah and Collins 1988) salivosa comb. nov., Int. J. Syst. Bacteriol., 42, 434, 1992.

577 16. Isogai, H. et al., Ecology of genus Porphyromonas in canine periodontal disease, Zentralbl. Veterinarmed. B, 46, 467, 1999. 17. Norris, J.M., and Love, D.N., Associations amongst three feline Porphyromonas species from the gingival margin of cats during periodontal health and disease, Vet. Microbiol., 65, 195, 1999. 18. Norris, J.M., and Love, D.N., The isolation and enumeration of three feline oral Porphyromonas species from subcutaneous abscesses in cats, Vet. Microbiol., 65, 115, 1999. 19. Allaker, R.P. et al., Prevalence of Porphyromonas and Prevotella species in the dental plaque of dogs, Vet. Rec., 140, 147, 1997. 20. Hirasawa, M., and Takada, K., Porphyromonas gingivicanis sp. nov. and Porphyromonas crevioricanis sp. nov., isolated from beagles, Int. J. Syst. Bacteriol., 44, 637, 1994. 21. Fournier, D. et al., Porphyromonas gulae sp. nov., an anaerobic, Gram-negative coccobacillus from the gingival sulcus of various animal hosts, Int. J. Syst. Evol. Microbiol., 51, 1179, 2001. 22. Mikkelsen, D. et al., Phylogenetic analysis of Porphyromonas species isolated from the oral cavity of Australian marsupials, Environ. Microbiol., 10, 2425, 2008. 23. Collins, J.G., Offenbacher, S., and Arnold, R.R., Effects of a combination therapy to eliminate Porphyromonas gingivalis in refractory periodontitis, J. Periodontol., 64, 998, 1993. 24. Blum, S. et al., Isolation of Porphyromonas levii from vaginal samples from cows in herds negative for bovine necrotic vulvovaginitis, Vet. Rec., 163, 745, 2008. 25. Finegold, S.M., and Jousimies-Somer, H., Recently described clinically important anaerobic bacteria: Medical aspects, Clin. Infect. Dis., 25 Suppl 2, 88, 1997. 26. Love, D.N., Porphyromonas macacae comb. nov., a consequence of Bacteroides macacae being a senior synonym of Polphyromonas salivosa, Int. J. Syst. Bacteriol., 45, 90, 1995. 27. Coykendall, A.L., Kaczmarek, F.S., and Slots, J., Genetic heterogeneity in Bacteroides asaccharolyticus (Holdeman and Moore, 1970; Finegold and Barnes, 1977; Approved Lists, 1980) and proposal of Bacteroides gingivalis sp. nov. and Bacteroides macacae (Slots and Genco) comb. nov., Int. J. Syst. Bacteriol., 30, 559, 1980. 28. Holt, J.G. et al., Group 6. Gram-negative, anaerobic, straight, curved, andhelical bacteria. In Bergey’s Manual of Determinative Bacteriology, 9th ed., pp. 291–334, The Williams & Wilkins Co., Baltimore, MD, 1994. 29. Mayrand, D., and Holt, S.C., Biology of asaccharolytic blackpigmented Bacteroides species, Microbiol. Rev., 52, 134, 1988. 30. Shah, H.N., Biochemical and chemical properties of Porphyromonas gingivalis. In Shah, H.N. (Editor), Biology of the Species Porphyromonas gingivalis, pp. 59–82, CRC Press, Boca Raton, 1993. 31. Shah, H.N. et al., Reclassification of Bacteroides levii (Holdeman, Cato & Moore) in the genus Porphyrornonas, as Porphyrornonas levii comb. nov., Int. J. Syst. Bacteriol., 45, 586, 1995. 32. Paster, B.J. et al., Phylogeny of Bacteroides, Prevotella and Porphyromonas spp. and related bacteria, J. Bacteriol., 176, 725, 1994. 33. Talan, D.A. et al., Bacteriologic analysis of infected dog and cat bites. Emergency Medicine Animal Bite Infection Study Group, N. Engl. J. Med., 340, 85, 1999. 34. de Lillo, A. et al., Culture-independent identification of periodontitis-associated Porphyromonas and Tannerella populations by targeted molecular analysis, J. Clin. Microbiol., 42, 5523, 2004.

578 35. Sayers, E.W. et al., Database resources of the National Center for Biotechnology Information, Nucleic Acids Res., 37, 5, 2009. 36. Benson, D.A. et al., GenBank, Nucleic Acids Res., 37, 26, 2009. 37. Hardham, J.M. et al., Transfer of Bacteroides splanchnicus to Odoribacter gen. nov. as Odoribacter splanchnicus comb. nov., and description of Odoribacter denticanis sp. nov., isolated from the crevicular spaces of canine periodontitis patients, Int. J. Syst. Evol. Microbiol., 58, 103, 2008. 38. Socransky, S.S., and Manganiello, S.D., The oral microbiota of man from birth to senility, J. Periodontol., 42, 485, 1971. 39. Salako, N.O. et al., The bacteriology of the supragingival plaque of child dental patients in Kuwait, Med. Princ. Pract., 13, 191, 2004. 40. Brook, I., Prevotella and Porphyromonas infections in children, J. Med. Microbiol., 42, 340, 1995. 41. Finegold, S.M., Anaerobic Bacteria in Human Disease, Academic Press, New York, 1977. 42. Dunkle, L.M., Brotherton, T.J., and Feigin, R.D., Anaerobic infections in children: A prospective study, Pediatrics, 57, 311, 1976. 43. Thirumoorthi, M.C., Keen, B.M., and Dajani, A.S., Anaerobic infections in children: A prospective survey, J. Clin. Microbiol., 3, 318, 1976. 44. Lopez, N.J., Occurrence of Actinobacillus actinomycetemcomitans, Porphyromonas gingivalis, and Prevotella intermedia in progressive adult periodontitis, J. Periodontol., 71, 948, 2000. 45. Ximenez-Fyvie, L.A. et al., Subgingival microbiota of periodontally untreated Mexican subjects with generalized aggressive periodontitis, J. Clin. Periodontol., 33, 869, 2006. 46. Herrera, D. et al., Subgingival microbial profiles in chronic periodontitis patients from Chile, Colombia and Spain, J. Clin. Periodontol., 35, 106, 2008. 47. Haffajee, A.D. et al., Subgingival microbiota of chronic periodontitis subjects from different geographic locations, J. Clin. Periodontol., 31, 996, 2004. 48. Sanz, M. et al., Differences in the composition of the subgingival microbiota of two periodontitis populations of different geographical origin. A comparison between Spain and The Netherlands, Eur. J. Oral Sci., 108, 383, 2000. 49. Rocas, I.N. et al., Prevalence of selected bacterial named species and uncultivated phylotypes in endodontic abscesses from two geographic locations, J. Endod., 32, 1135, 2006. 50. Foschi, F. et al., Detection of bacteria in endodontic samples by polymerase chain reaction assays and association with defined clinical signs in Italian patients, Oral Microbiol. Immunol., 20, 289, 2005. 51. Lopez, N.J. et al., Subgingival microbiota of chilean patients with chronic periodontitis, J. Periodontol., 75, 717, 2004. 52. Machado de Oliveira, J.C. et al., Bacterial community profiles of endodontic abscesses from Brazilian and USA subjects as compared by denaturing gradient gel electrophoresis analysis, Oral Microbiol. Immunol., 22, 14, 2007. 53. Baumgartner, J.C. et al., Geographical differences in bacteria detected in endodontic infections using polymerase chain reaction, J. Endod., 30, 141, 2004. 54. Siqueira, J.F. et al., Differences in prevalence of selected bacterial species in primary endodontic infections from two distinct geographic locations, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 99, 641, 2005. 55. Kim, S.A., and Lim, S.S., T lymphocyte subpopulations and interleukin-2, interferon-gamma, and interleukin-4 in rat pulpitis experimentally induced by specific bacteria, J. Endod., 28, 202, 2002.

Molecular Detection of Human Bacterial Pathogens 56. Lemke, J.L. et al., Description of freshwater bacterial assemblages from the upper Paraná river floodpulse system, Brazil, Microb. Ecol., 57, 94, 2009. 57. Jansen, H.J. et al., Degradation of immunoglobulin G by periodontal bacteria, Oral Microbiol. Immunol., 9, 345, 1994. 58. Magalashvili, L. et al., Cytokine release and expression induced by OmpA proteins from the Gram-negative anaerobes, Porphyromonas asaccharolytica and Bacteroides fragilis, FEMS Immunol. Med. Microbiol., 53, 252, 2008. 59. Hohne, C. et al., Occurrence of plasmids in black-pigmented Gram-negative anaerobes, FEMS Immunol. Med. Microbiol., 6, 103, 1993. 60. Odell, E.W., and Wu, P.J., Susceptibility of Porphyromonas gingivalis and P. asaccharolytica to the nonoxidative killing mechanisms of human neutrophils, Arch. Oral Biol., 37, 597, 1992. 61. Bahar, H. et al., Antimicrobial resistance and beta-lactamase production of clinical isolates of Prevotella and Porphyromonas species, Chemotherapy, 51, 9, 2005. 62. Egwari, L. et al., A study of the anaerobic bacterial flora of the female genital tract in health and disease, Cent. Afr. J. Med., 41, 391, 1995. 63. Holst, E., Goffeng, A.R., and Andersch, B., Bacterial vaginosis and vaginal microorganisms in idiopathic premature labor and association with pregnancy outcome, J. Clin. Microbiol., 32, 176, 1994. 64. Duerden, B.I., Black-pigmented Gram-negative anaerobes in genito-urinary tract and pelvic infections, FEMS Immunol. Med. Microbiol., 6, 223, 1993. 65. Neut, C. et al., Treatment of diversion colitis with short-chain fatty acids. Bacteriological study, Gastroenterol. Clin. Biol., 19, 871, 1995. 66. Sharma, D., and Rouphael, N., Case report: Atypical Lemierre’s disease secondary to Porphyromonas asaccharolytica, Curr. Opin. Pediatr., 21, 397, 2009. 67. Brook, I., and Frazier, E.H., Bacteriology of chronic maxillary sinusitis associated with nasal polyposis, J. Med. Microbiol., 54, 595, 2005. 68. Castagnola, E. et al., Broviac catheter-related bacteraemias due to unusual pathogens in children with cancer: Case reports with literature review, J. Infect., 34, 215, 1997. 69. Revankar, S.G., and Clark, R.A., Infected cardiac myxoma. Case report and literature review, Medicine (Baltimore), 77, 337, 1998. 70. Tran, T. et al., Absence of Porphyromonas asaccharolytica, Bacteroides fragilis and Chlamydia pneumoniae in human subgingival plaque, Oral Microbiol. Immunol., 12, 377, 1997. 71. Hall, E.R. et al., The gingival immune response to Actinobacillus actinomycetemcomitans in juvenile periodontitis, J. Periodontol., 62, 792, 1991. 72. Rawlinson, A., Duerden, B.I., and Goodwin, L., New findings on the microbial flora associated with adult periodontitis, J. Dent., 21, 179, 1993. 73. Ueno, K. et al., Cytotoxicity of anaerobic bacteria isolated from infected root canal, Kokubyo Gakkai Zasshi, 60, 408, 1993. 74. Peters, L.B. et al., Viable bacteria in root dentinal tubules of teeth with apical periodontitis, J. Endod., 27, 76, 2001. 75. Rawlinson, A., Duerden, B.I., and Goodwin, L., Effects of root planing on the distribution of microorganisms adult periodontitis sites, Eur. J. Prosthodont. Restor. Dent, 1, 103, 1993. 76. Moore, L.V., and Moore, W.E., Oribaculum catoniae gen. nov., sp. nov.; Catonella morbi gen. nov., sp. nov.; Hallella seregens gen. nov., sp. nov.; Johnsonella ignava gen. nov., sp.

Porphyromonas









nov.; and Dialister pneumosintes gen. nov., comb. nov., nom. rev., Anaerobic Gram-negative bacilli from the human gingival crevice, Int. J. Syst. Bacteriol., 44, 187, 1994. 77. Moore, W.E. et al., Comparative bacteriology of juvenile periodontitis, Infect. Immun., 48, 507, 1985. 78. Moore, W.E. et al., The microflora of periodontal sites showing active destructive progression, J. Clin. Periodontol., 18, 729, 1991. 79. Zerr, M.A. et al., Growth stimulation of Porphyromonas endodontalis by hemoglobin and protoporphyrin IX, Oral Microbiol. Immunol., 15, 365, 2000. 80. Persson, S. et al., The formation of hydrogen sulfide and methyl mercaptan by oral bacteria, Oral Microbiol. Immunol., 5, 195, 1990. 81. Ho, Y.C., and Chang, Y.C., Effects of a bacterial lipid byproduct on human pulp fibroblasts in vitro, J. Endod., 33, 437, 2007. 82. Rocas, I.N., and Siqueira, J.F.Jr., Root canal microbiota of teeth with chronic apical periodontitis, J. Clin. Microbiol., 46, 3599, 2008. 83. Sundqvist, G., Associations between microbial species in dental root canal infections, Oral Microbiol. Immunol., 7, 257, 1992. 84. Zerr, M. et al., Porphyromonas endodontalis binds, reduces and grows on human hemoglobin, Oral Microbiol. Immunol., 16, 229, 2001. 85. Rosen, G. et al., The purification and characterization of an 88-kDa Porphyromonas endodontalis 35406 protease, Oral Microbiol. Immunol., 16, 326, 2001. 86. Horiba, N. et al., Complement activation by lipopolysaccharides purified from Gram-negative bacteria isolated from infected root canals, Oral Surg. Oral Med. Oral Pathol., 74, 648, 1992. 87. Sundqvist, G. et al., Degradation of human immunoglobulins G and M and complement factors C3 and C5 by blackpigmented Bacteroides, J. Med. Microbiol., 19, 85, 1985. 88. Chang, J. et al., NF-kappaB activation in human dental pulp stem cells by TNF and LPS, J. Dent. Res., 84, 994, 2005. 89. Yang, L.C. et al., Induction of interleukin-8 gene expression by black-pigmented Bacteroides in human pulp fibroblasts and osteoblasts, Int. Endod. J., 36, 774, 2003. 90. Chen, S.C. et al., The upregulation of receptor activator NF-kappaB ligand expression by interleukin-1alpha and Porphyromonas endodontalis in human osteoblastic cells, Int. Endod. J., 42, 375, 2009. 91. Hosoya, S. et al., Stimulation of interleukin-1beta-independent interleukin-6 production in human dental pulp cells by lipopolysaccharide, Biochem. Mol. Med., 59, 138, 1996. 92. Ko, H.J., and Lim, S.S., Production of macrophage inflammatory protein (MIP)-1alpha and MIP-1beta by human polymorphonuclear neutrophils stimulated with Porphyromonas endodontalis lipopolysaccharide, J. Endod., 28, 754, 2002. 93. Hong, C.Y. et al., The role of lipopolysaccharide in infectious bone resorption of periapical lesion, J. Oral Pathol. Med., 33, 162, 2004. 94. Chang, Y.C. et al., Stimulation of matrix metalloproteinases by black-pigmented Bacteroides in human pulp and periodontal ligament cell cultures, J. Endod., 28, 90, 2002. 95. Yang, L.C. et al., Induction of vascular endothelial growth factor expression in human pulp fibroblasts stimulated with black-pigmented Bacteroides, Int. Endod. J., 37, 588, 2004. 96. Jiang, Y., and Schilder, H., An optimal host response to a bacterium may require the interaction of leukocytes and resident host cells, J. Endod., 28, 279, 2002.

579 97. Chang, Y.C. et al., Induction of cyclooxygenase-2 mRNA and protein expression in human pulp cells stimulated with blackpigmented Bacteroides, J. Endod., 29, 240, 2003. 98. Huang, F.M. et al., Examination of the signal transduction pathways leading to upregulation of tissue type plasminogen activator by Porphyromonas endodontalis in human pulp cells, Int. Endod. J., 38, 860, 2005. 99. Yang, S.F. et al., Effect of black-pigmented bacteria on the plasminogen-plasmin system in human pulp and osteoblastic cells, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 95, 621, 2003. 100. Dorn, B.R. et al., Invasion of vascular cells in vitro by Porphyromonas endodontalis, Int. Endod. J., 35, 366, 2002. 101. Siqueira, J.F. Jr. et al., Molecular detection of black-pigmented bacteria in infections of endodontic origin, J. Endod., 27, 563, 2001. 102. Blome, B. et al., Molecular identification and quantification of bacteria from endodontic infections using real-time polymerase chain reaction, Oral Microbiol. Immunol., 23, 384, 2008. 103. Sunde, P.T. et al., Assessment of periradicular microbiota by DNA–DNA hybridization, Endod. Dent. Traumatol., 16, 191, 2000. 104. Bogen, G., and Slots, J., Black-pigmented anaerobic rods in closed periapical lesions, Int. Endod. J., 32, 204, 1999. 105. Vickerman, M.M. et al., Phylogenetic analysis of bacterial and archaeal species in symptomatic and asymptomatic endodontic infections, J. Med. Microbiol., 56, 110, 2007. 106. Tomazinho, L.F., and Avila-Campos, M.J., Detection of Porphyromonas gingivalis, Porphyromonas endodontalis, Prevotella intermedia, and Prevotella nigrescens in chronic endodontic infection, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 103, 285, 2007. 107. Siqueira, J.F. Jr. et al., Bacteria in the apical root canal of teeth with primary apical periodontitis, Oral Surg. Oral Med. Oral Pathol. Oral Radiol Endod, 107, 721, 2009. 108. Gomes, B.P. et al., Porphyromonas gingivalis, Porphyromonas endodontalis, Prevotella intermedia and Prevotella nigrescens in endodontic lesions detected by culture and by PCR, Oral Microbiol. Immunol., 20, 211, 2005. 109. Martin, F.E. et al., Quantitative microbiological study of human carious dentine by culture and real-time PCR: Association of anaerobes with histopathological changes in chronic pulpitis, J. Clin. Microbiol., 40, 1698, 2002. 110. Persson, G.R. et al., Tannerella forsythia and Pseudomonas aeruginosa in subgingival bacterial samples from parous women, J. Periodontol., 79, 508, 2008. 111. Kumar, P.S. et al., New bacterial species associated with chronic periodontitis, J. Dent. Res., 82, 338, 2003. 112. van Winkelhoff, A.J., van Steenbergen, T.J., and de Graaff, J., Porphyromonas (Bacteroides) endodontalis: Its role in endodontal infections, J. Endod., 18, 431, 1992. 113. Enersen, M. et al., fimA genotypes and multilocus sequence types of Porphyromonas gingivalis from patients with periodontitis, J. Clin. Microbiol., 46, 31, 2008. 114. Hamada, S. et al., The importance of fimbriae in the virulence and ecology of some oral bacteria, Oral Microbiol. Immunol., 13, 129, 1998. 115. Suzuki, N., Nakano, Y., and Kiyoura, Y., Characterizing the specific coaggregation between Actinobacillus actinomycetemcomitans serotype c strains and Porphyromonas gingivalis ATCC 33277, Oral Microbiol. Immunol., 21, 385, 2006. 116. Yamada, M., Ikegami, A., and Kuramitsu, H.K., Synergistic biofilm formation by Treponema denticola and Porphyromonas gingivalis, FEMS Microbiol. Lett., 250, 271, 2005.

580 117. Maeda, K. et al., Oral streptococcal glyceraldehyde-3-phosphate dehydrogenase mediates interaction with Porphyromonas gingivalis fimbriae, Microbes. Infect., 6, 1163, 2004. 118. Hongo, H., Takano, H., and Morita, M., Dense fimbrial meshwork enhances Porphyromonas gingivalis adhesiveness: A scanning electron microscopic study, J. Periodontal. Res., 42, 114, 2007. 119. Ogawa, T., and Hamada, S., Hemagglutinating and chemotactic properties of synthetic peptide segments of fimbrial protein from Porphyromonas gingivalis, Infect. Immun., 62, 3305, 1994. 120. Lewis, J.P. et al., Hemoglobinase activity of the lysine gingipain protease (Kgp) of Porphyromonas gingivalis W83, J. Bacteriol., 181, 4905, 1999. 121. Lewis, J.P., and Macrina, F.L., Localization of HArepcontaining genes on the chromosome of Porphyromonas gingivalis W83, Infect. Immun., 67, 2619, 1999. 122. Slaney, J.M. et al., Mechanisms of resistance of Porphyromonas gingivalis to killing by serum complement, Infect. Immun., 74, 5352, 2006. 123. Liu, R. et al., P. gingivalis and E. coli lipopolysaccharides exhibit different systemic but similar local induction of inflammatory markers, J. Periodontol. 79, 1241, 2008. 124. Domon, H. et al., Up-regulation of the endoplasmic reticulum stress-response in periodontal disease, Clin. Chim. Acta., 401, 134, 2009. 125. Hajishengallis, G., Porphyromonas gingivalis-host interactions: Open war or intelligent guerilla tactics? Microbes. Infect., 11, 637, 2009. 126. Ebersole, J.L. et al., Comparative virulence of periodontopathogens in a mouse abscess model, Oral Dis., 1, 115, 1995. 127. Yilmaz, O., The chronicles of Porphyromonas gingivalis: The microbium, the human oral epithelium and their interplay, Microbiology, 154, 2897, 2008. 128. Hajishengallis, G. et al., Subversion of innate immunity by periodontopathic bacteria via exploitation of complement receptor-3, Adv. Exp. Med. Biol., 632, 203, 2008. 129. Yoneda, M. et al., Mixed infection of Porphyromonas gingivalis and Bacteroides forsythus in a murine abscess model: Involvement of gingipains in a synergistic effect, J. Periodontal. Res., 36, 237, 2001. 130. Diaz, P.I. et al., Molecular characterization of subject-specific oral microflora during initial colonization of enamel, Appl. Environ. Microbiol., 72, 2837, 2006. 131. Amano, A. et al., Variations of Porphyromonas gingivalis fimbriae in relation to microbial pathogenesis, J. Periodontal. Res., 39, 136, 2004. 132. Njoroge, T. et al., A role for fimbriae in Porphyromonas gingivalis invasion of oral epithelial cells, Infect. Immun., 65, 1980, 1997. 133. Okuda, K., Slots, J., and Genco, J., Bacteroides gingivalis, Bacteroides asaccharolyticus and Bacteroides melanogenicus subspecies: Cell surface morphology and adherence to erythrocytes and human buccal epithelial cells, Curr. Microbiol., 6, 7, 1981. 134. Yoshimura, F. et al., Fimbriae from the oral anaerobe Bacteroides gingivalis: Physical, chemical, and immunological properties, J. Bacteriol., 163, 730, 1985. 135. Takahashi, Y. et al., Detection of fimbrilin gene (fimA) in Porphyromonas (Bacteroides) gingivalis by Southern blot analysis, J. Periodontal. Res., 27, 599, 1992. 136. Gibson, F.C., and Genco, C.A., The Genus Porphyromonas. In Dworkin, M., Falkow, S., Rosenberg, E., Schleifer, K.H., and Stackebrandt, E. (Editors), The Prokaryotes: A Handbook

Molecular Detection of Human Bacterial Pathogens on the Biology of Bacteria, pp. 428–454, Springer, Singapore, 2006. 137. Olsen, I., Shah, H.N., and Gharbia, S.E., Taxonomy and biochemical characteristics of Actinobacillus actinomycetemcomitans and Porphyromonas gingivalis, Periodontol. 2000, 20, 14, 1999. 138. Kontani, M. et al., Adherence of Porphyromonas gingivalis to matrix proteins via a fimbrial cryptic receptor exposed by its own arginine-specific protease, Mol. Microbiol., 24, 1179, 1997. 139. Amano, A. et al., Structural domains of Porphyromonas gingivalis recombinant fimbrillin that mediate binding to salivary proline-rich protein and statherin, Infect. Immun., 64, 1631, 1996. 140. Amano, A. et al., Binding of Porphyromonas gingivalis fimbriae to proline-rich glycoproteins in parotid saliva via a domain shared by major salivary components, Infect. Immun., 66, 2072, 1998. 141. Goulbourne, P.A., and Ellen, R.P., Evidence that Porphyromonas (Bacteroides) gingivalis fimbriae function in adhesion to Actinomyces viscosus, J. Bacteriol., 173, 5266, 1991. 142. Lamont, R.J. et al., Involvement of Porphyromonas gingivalis fimbriae in adherence to Streptococcus gordonii, Oral Microbiol. Immunol., 8, 272, 1993. 143. Murakami, Y. et al., Porphyromonas gingivalis fimbrillin is one of the fibronectin-binding proteins, Infect. Immun., 64, 2571, 1996. 144. Sojar, H.T., Hamada, N., and Genco, R.J., Structures involved in the interaction of Porphyromonas gingivalis fimbriae and human lactoferrin, FEBS Lett., 422, 205, 1998. 145. Umemoto, T., and Hamada, N., Characterization of biologically active cell surface components of a periodontal pathogen. The roles of major and minor fimbriae of Porphyromonas gingivalis, J. Periodontol., 74, 119, 2003. 146. Hu, S.W. et al., Transvascular dissemination of Porphyromonas gingivalis from a sequestered site is dependent upon activation of the kallikrein/kinin pathway, J. Periodontal. Res., 41, 200, 2006. 147. Imamura, T., The role of gingipains in the pathogenesis of periodontal disease, J. Periodontol., 74, 111, 2003. 148. Wingrove, J.A. et al., Activation of complement components C3 and C5 by a cysteine proteinase (gingipain-1) from Porphyromonas (Bacteroides) gingivalis, J. Biol. Chem., 267, 18902, 1992. 149. Cutler, C.W., Arnold, R.R., and Schenkein, H.A., Inhibition of C3 and IgG proteolysis enhances phagocytosis of Porphyromonas gingivalis, J. Immunol., 151, 7016, 1993. 150. Hawlisch, H. et al., C5a negatively regulates toll-like receptor 4-induced immune responses, Immunity, 22, 415, 2005. 151. Kumagai, Y. et al., Molecular mechanism for connective tissue destruction by dipeptidyl aminopeptidase IV produced by the periodontal pathogen Porphyromonas gingivalis, Infect. Immun., 73, 2655, 2005. 152. Holt, S.C. et al., Virulence factors of Porphyromonas gingivalis, Periodontol. 2000, 20, 168, 1999. 153. Potempa, J., Banbula, A., and Travis, J., Role of bacterial proteinases in matrix destruction and modulation of host responses, Periodontol. 2000, 24, 153, 2000. 154. Sroka, A. et al., Degradation of host heme proteins by lysineand arginine-specific cysteine proteinases (gingipains) of Porphyromonas gingivalis, J. Bacteriol., 183, 5609, 2001. 155. Liu, X. et al., Coordinate expression of the Porphyromonas gingivalis lysine-specific gingipain proteinase, Kgp,

Porphyromonas arginine-specific gingipain proteinase, RgpA, and the heme/ hemoglobin receptor, HmuR, Biol. Chem., 385, 1049, 2004. 156. Wojtowicz, H. et al., Unique structure and stability of HmuY, a novel heme-binding protein of Porphyromonas gingivalis, PLoS Pathog., 5, e1000419, 2009. 157. Simpson, W., Olczak, T., and Genco, C.A., Characterization and expression of HmuR, a TonB-dependent hemoglobin receptor of Porphyromonas gingivalis, J. Bacteriol., 182, 5737, 2000. 158. Darveau, R.P. et al., Ability of bacteria associated with chronic inflammatory disease to stimulate E-selectin expression and promote neutrophil adhesion, Infect. Immun., 63, 1311, 1995. 159. Darveau, R.P. et al., Porphyromonas gingivalis lipopolysaccharide contains multiple lipid A species that functionally interact with both toll-like receptors 2 and 4, Infect. Immun., 72, 5041, 204. 160. Al-Qutub, M.N. et al., Hemin-dependent modulation of the lipid A structure of Porphyromonas gingivalis lipopolysaccharide, Infect. Immun., 74, 4474, 2006. 161. Lamont, R.J., and Jenkinson, H.F., Life below the gum line: Pathogenic mechanisms of Porphyromonas gingivalis, Microbiol. Mol. Biol. Rev., 62, 1244, 1998. 162. Domon, H. et al., Early and preferential induction of IL-1 receptor-associated kinase-M in THP-1 cells by LPS derived from Porphyromonas gingivalis, J. Leukoc. Biol., 83, 672, 2008. 163. Cunningham, M.D. et al., Helicobacter pylori and Porphyromonas gingivalis lipopolysaccharides are poorly transferred to recombinant soluble CD14, Infect. Immun., 64, 3601, 1996. 164. Isogai, E. et al., In vivo induction of apoptosis and immune responses in mice by administration of lipopolysaccharide from Porphyromonas gingivalis, Infect. Immun., 64, 1461, 1996. 165. Haapasalo, M., Kerosuo, E., and Lounatmaa, K., Hydrophobicities of human polymorphonuclear leukocytes and oral Bacteroides and Porphyromonas spp., Wolinella recta, and Eubacterium yurii with special reference to bacterial surface structures, Scand. J. Dent. Res., 98, 472, 1990. 166. Ingham, H.R. et al., Inhibition of phagocytosis in vitro by obligate anaerobes, Lancet, 2, 1252, 1977. 167. Laine, M.L., and van Winkelhoff, A.J., Virulence of six capsular serotypes of Porphyromonas gingivalis in a mouse model, Oral Microbiol. Immunol., 13, 322, 1998. 168. Laine, M.L., Appelmelk, B.J., and van Winkelhoff, A.J., Prevalence and distribution of six capsular serotypes of Porphyromonas gingivalis in periodontitis patients, J. Dent. Res., 76, 1840, 1997. 169. Laine, M.L., Appelmelk, B.J., and van Winkelhoff, A.J., Novel polysaccharide capsular serotypes in Porphyromonas gingivalis, J. Periodontal. Res., 31, 278, 1996. 170. Furuta, N., Takeuchi, H., and Amano, A., Entry of Porphyromonas gingivalis outer membrane vesicles into epithelial cells causes cellular functional impairment, Infect. Immun., 77, 4761, 2009. 171. Furuta, N. et al., Porphyromonas gingivalis outer membrane vesicles enter human epithelial cells via an endocytic pathway and are sorted to lysosomal compartments, Infect. Immun., 77, 4187, 2009. 172. Kurita-Ochiai, T., Fukushima, K., and Ochiai, K., Butyric acid-induced apoptosis of murine thymocytes, splenic T cells, and human Jurkat T cells, Infect. Immun., 65, 35, 1997. 173. Kurita-Ochiai, T., Ochiai, K., and Fukushima, K., Volatile fatty acid, metabolic by-product of periodontopathic bacteria,

581 induces apoptosis in WEHI 231 and RAJI B lymphoma cells and splenic B cells, Infect. Immun., 66, 2587, 1998. 174. Borrell, L.N., and Papapanou, P.N., Analytical epidemiology of periodontitis, J. Clin. Periodontol., 32, 132, 2005. 175. Tanner, A. et al., Microbiota of health, gingivitis, and initial periodontitis, J. Clin. Periodontol., 25, 85, 1998. 176. Cortelli, J.R. et al., Detection of periodontal pathogens in oral mucous membranes of edentulous individuals, J. Periodontol., 79, 1962, 2008. 177. Cortelli, J.R. et al., Etiological analysis of initial colonization of periodontal pathogens in oral cavity, J. Clin. Microbiol., 46, 1322, 2008. 178. Ximenez-Fyvie, L.A., Haffajee, A.D., and Socransky, S.S., Comparison of the microbiota of supra- and subgingival plaque in health and periodontitis, J. Clin. Periodontol., 27, 648, 2000. 179. Haffajee, A.D. et al., Subgingival microbiota in healthy, well-maintained elder and periodontitis subjects, J. Clin. Periodontol., 25, 346, 1998. 180. Socransky, S.S. et al., Microbial complexes in subgingival plaque, J. Clin. Periodontol., 25, 134, 1998. 181. Colombo, A.P. et al., Clinical and microbiological features of refractory periodontitis subjects, J. Clin. Periodontol., 25, 169, 1998. 182. Loos, B.G. et al., Genetic structure of populations of Porphyromonas gingivalis associated with periodontitis and other oral infections, Infect. Immun., 61, 204, 1993. 183. Mager, D.L. et al., Distribution of selected bacterial species on intraoral surfaces, J. Clin. Periodontol., 30, 644, 2003. 184. Beikler, T. et al., Sampling strategy for intraoral detection of periodontal pathogens before and following periodontal therapy, J. Periodontol., 77, 1323, 2006. 185. Gaetti-Jardim, E. Jr. et al., Quantitative detection of periodontopathic bacteria in atherosclerotic plaques from coronary arteries, J. Med. Microbiol., 58, 1568, 2009. 186. Rudolph, T. et al., 16S rDNA PCR analysis of infectious keratitis: A case series, Acta. Ophthalmol. Scand., 82, 463, 2004. 187. Perez-Chaparro, P.J. et al., Genotypic characterization of Porphyromonas gingivalis isolated from subgingival plaque and blood sample in positive bacteremia subjects with periodontitis, J. Clin. Periodontol., 35, 748, 2008. 188. Iida, Y. et al., Brain abscess in which Porphyromonas gingivalis was detected in cerebrospinal fluid, Br. J. Oral Maxillofac. Surg., 42, 180, 2004. 189. Hirata, R. Jr. et al., Isolation of Porphyromonas gingivalis strain from tubal-ovarian abscess, J. Clin. Microbiol., 33, 1925, 1995. 190. Soskolne, W.A., and Klinger, A., The relationship between periodontal diseases and diabetes: An overview, Ann. Periodontol., 6, 91, 2001. 191. Matthews, D.C., The relationship between diabetes and periodontal disease, J. Can. Dent. Assoc., 68, 161, 2002. 192. Seymour, G.J. et al., Infection or inflammation: The link between periodontal and cardiovascular diseases, Future Cardiol., 5, 5, 2009. 193. Seymour, G.J. et al., Relationship between periodontal infections and systemic disease, Clin. Microbiol. Infect., 13 Suppl 4, 3, 2007. 194. Jousimies-Somer, H.R., Update on the taxonomy and the clinical and laboratory characteristics of pigmented anaerobic Gram-negative rods, Clin. Infect. Dis., 20 Suppl 2, 187, 1995. 195. Stoner, K.A. et al., Quantitative survival of aerobic and anaerobic microorganisms in Port-A-Cul and Copan transport systems, J. Clin. Microbiol., 46, 2739, 2008.

582 196. van Steenbergen, T.J. et al., Survival in transport media of Actinobacillus actinomycetemcomitans, Porphyromonas gingivalis and Prevotella intermedia in human subgingival samples, Oral Microbiol. Immunol., 8, 370, 1993. 197. Falagas, M.E., and Siakavellas, E., Bacteroides, Prevotella, and Porphyromonas species: A review of antibiotic resistance and therapeutic options, Int. J. Antimicrob. Agents, 15, 1, 2000. 198. Stingu, C.S. et al., Rapid identification of oral anaerobic bacteria cultivated from subgingival biofilm by MALDI-TOF-MS, Oral Microbiol. Immunol., 23, 372, 2008. 199. Kitch, T.T., and Appelbaum, P.C., Accuracy and reproducibility of the 4-hour ATB 32A method for anaerobe identification, J. Clin. Microbiol., 27, 2509, 1989. 200. Assmus, B. et al., Direct examination of subgingival plaque from a diseased periodontal site using confocal laser scanning microscopy (CLSM), New Microbiol., 20, 155, 1997. 201. Wolff, L.F. et al., Fluorescence immunoassay for detecting periodontal bacterial pathogens in plaque, J. Clin. Microbiol., 29, 1645, 1991. 202. van der Ploeg, J.R. et al., Quantitative detection of Porphyromonas gingivalis fimA genotypes in dental plaque, FEMS Microbiol. Lett., 232, 31, 2004. 203. Pischon, N. et al., Effects of Porphyromonas gingivalis on cell cycle progression and apoptosis of primary human chondrocytes, Ann. Rheum. Dis., 68, 1902, 2009. 204. Friedrichs, C. et al., Rapid identification of viridans streptococci by mass spectrometric discrimination, J. Clin. Microbiol., 45, 2392, 2007. 205. Ruelle, V. et al., Rapid identification of environmental bacterial strains by matrix-assisted laser desorption/ionization timeof-flight mass spectrometry, Rapid Commun. Mass Spectrom., 18, 2013, 2004. 206. Fagerquist, C.K. et al., Sub-speciating Campylobacter jejuni by proteomic analysis of its protein biomarkers and their post-translational modifications, J. Proteome Res., 5, 2527, 2006. 207. Rupf, S. et al., Differentiation of mutans streptococci by intact cell matrix-assisted laser desorption/ionization time-of-flight mass spectrometry, Oral Microbiol. Immunol., 20, 267, 2005. 208. Loesche, W.J. et al., Development of a diagnostic test for anaerobic periodontal infections based on plaque hydrolysis of benzoyl-DL-arginine-naphthylamide, J. Clin. Microbiol., 28, 1551, 1990. 209. Gemmell, E. et al., Antibody responses of Porphyromonas gingivalis infected gingivitis and periodontitis subjects, Oral Dis., 1, 63, 1995. 210. Teanpaisan, R., and Douglas, C.W., Molecular fingerprinting of Porphyromonas gingivalis by PCR of repetitive extragenic palindromic (REP) sequences and comparison with other fingerprinting methods, J. Med. Microbiol., 48, 741, 1999. 211. Boutaga, K. et al., Comparison of real-time PCR and culture for detection of Porphyromonas gingivalis in subgingival plaque samples, J. Clin. Microbiol., 41, 4950, 2003. 212. 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. 213. Rupf, S. et al., Comparison of profiles of key periodontal pathogens in periodontium and endodontium, Endod. Dent. Traumatol., 16, 269, 2000. 214. Morillo, J.M. et al., Quantitative real-time polymerase chain reaction based on single copy gene sequence for detection of periodontal pathogens, J. Clin. Periodontol., 31, 1054, 2004.

Molecular Detection of Human Bacterial Pathogens 215. Doungudomdacha, S., Rawlinson, A., and Douglas, C.W., Enumeration of Porphyromonas gingivalis, Prevotella intermedia and Actinobacillus actinomycetemcomitans in subgingival plaque samples by a quantitative-competitive PCR method, J. Med. Microbiol., 49, 861, 2000. 216. Sanz, M. et al., Methods of detection of Actinobacillus actinomycetemcomitans, Porphyromonas gingivalis and Tannerella forsythensis in periodontal microbiology, with special emphasis on advanced molecular techniques: A review, J. Clin. Periodontol., 31, 1034, 2004. 217. Sakamoto, M., Umeda, M., and Benno, Y., Molecular analysis of human oral microbiota, J. Periodontal. Res., 40, 277, 2005. 218. Rupf, S., Merte, K., and Eschrich, K., Quantification of bacteria in oral samples by competitive polymerase chain reaction, J. Dent. Res., 78, 850, 1999. 219. Saito, D. et al., Real-time polymerase chain reaction quantification of Porphyromonas gingivalis and Tannerella forsythia in primary endodontic infections, J. Endod., 35, 1518, 2009. 220. Slots, J. et al., Detection of putative periodontal pathogens in subgingival specimens by 16S ribosomal DNA amplification with the polymerase chain reaction, Clin. Infect. Dis., 20 Suppl 2, 304, 1995. 221. Tran, S.D., and Rudney, J.D., Improved multiplex PCR using conserved and species-specific 16S rRNA gene primers for simultaneous detection of Actinobacillus actinomycetemcomitans, Bacteroides forsythus, and Porphyromonas gingivalis, J. Clin. Microbiol., 37, 3504, 1999. 222. Haffajee, A.D. et al., Comparison between polymerase chain reaction-based and checkerboard DNA hybridization techniques for microbial assessment of subgingival plaque samples, J. Clin. Periodontol., 36, 642, 2009. 223. Verner, C. et al., Carpegen real-time polymerase chain reaction vs. anaerobic culture for periodontal pathogen identification, Oral Microbiol. Immunol., 21, 341, 2006. 224. Menard, C., and Mouton, C., Clonal diversity of the taxon Porphyromonas gingivalis assessed by random amplified polymorphic DNA fingerprinting, Infect. Immun., 63, 2522, 1995. 225. Luterbacher, S. et al., Diagnostic characteristics of clinical and microbiological tests for monitoring periodontal and periimplant mucosal tissue conditions during supportive periodontal therapy (SPT), Clin. Oral Implants Res., 11, 521, 2000. 226. Eguchi, T. et al., Microbial changes in patients with acute periodontal abscess after treatment detected by PadoTest, Oral Dis., 14, 180, 2008. 227. Socransky, S.S. et al., “Checkerboard” DNA–DNA hybridization, Biotechniques, 17, 788, 1994. 228. Paster, B.J., Bartoszyk, I.M., and Dewhirst, F.E., Identification of oral streptococci using PCR-based, reverse-capture, checkerboard hybridization, Methods Cell Sci., 20, 223, 1998. 229. Koehler, A. et al., Multilocus sequence analysis of Porphyromonas gingivalis indicates frequent recombination, Microbiology, 149, 2407, 2003. 230. Colombo, A.P. et al., Comparisons of subgingival microbial profiles of refractory periodontitis, severe periodontitis, and periodontal health using the human oral microbe identification microarray, J. Periodontol., 80, 1421, 2009. 231. Eberhard, J. et al., The stage of native biofilm formation determines the gene expression of human beta-defensin-2, psoriasin, ribonuclease 7 and inflammatory mediators: A novel approach for stimulation of keratinocytes with in situ formed biofilms, Oral Microbiol. Immunol., 23, 21, 2008. 232. Zijnge, V. et al., Denaturing gradient gel electrophoresis as a diagnostic tool in periodontal microbiology, J. Clin. Microbiol., 44, 3628, 2006.

Porphyromonas 233. Takeshita, T. et al., The ecological proportion of indigenous bacterial populations in saliva is correlated with oral health status, ISME J., 3, 65, 2009. 234. Boutaga, K. et al., Comparison of subgingival bacterial sampling with oral lavage for detection and quantification of periodontal pathogens by real-time polymerase chain reaction, J. Periodontol., 78, 79, 2007. 235. Krigar, D.M. et al., Two subgingival plaque-sampling strategies used with RNA probes, J. Periodontol., 78, 72, 2007. 236. Teles, F.R., Haffajee, A.D., and Socransky, S.S., The reproducibility of curet sampling of subgingival biofilms, J. Periodontol., 79, 705, 2008. 237. Jervoe-Storm, P.M. et al., Comparison of curet and paper point sampling of subgingival bacteria as analyzed by realtime polymerase chain reaction, J. Periodontol., 78, 909, 2007. 238. Al-Robaiy, S., Rupf, S., and Eschrich, K., Rapid competitive PCR using melting curve analysis for DNA quantification, Biotechniques, 31, 1382, 1388, 2001. 239. Rupf, S. et al., Comparison of different techniques of quantitative PCR for determination of Streptococcus mutans counts in saliva samples, Oral Microbiol. Immunol., 18, 50, 2003.

583 240. Rupf, S. et al., In vitro, clinical, and microbiological evaluation of a linear oscillating device for scaling and root planing, J. Periodontol., 76, 1942, 2005. 241. Rupf, S. et al., Quantitative determination of Streptococcus mutans by using competitive polymerase chain reaction, Eur. J. Oral Sci., 107, 75, 1999. 242. Paster, B.J. et al., The breadth of bacterial diversity in the human periodontal pocket and other oral sites, Periodontol. 2000, 42, 80, 2006. 243. Darveau, R.P., The oral microbial consortium’s interaction with the periodontal innate defense system, DNA Cell Biol., 28, 389, 2009. 244. Frandsen, E.V. et al., Evidence of recombination in Porphyromonas gingivalis and random distribution of putative virulence markers, Infect. Immun., 69, 4479, 2001. 245. Byrne, S.J. et al., Progression of chronic periodontitis can be predicted by the levels of Porphyromonas gingivalis and Treponema denticola in subgingival plaque, Oral Microbiol. Immuno.l, 24, 469, 2009. 246. Walter, C. et al., Critical assessment of microbiological diagnostics in periodontal diseases with special focus on Porphyromonas gingivalis, Schweiz. Monatsschr. Zahnmed., 115, 415, 2005.

51 Prevotella Mario J. Avila-Campos, Maria R.L. Simionato, and Elerson Gaetti-Jardim Jr. CONTENTS 51.1 Introduction...................................................................................................................................................................... 585 51.1.1 Classification, Morphology, and Biology................................................................................................... 585 51.1.2 Clinical Features............................................................................................................................................. 587 51.1.3 Virulence and Pathogenesis........................................................................................................................... 589 51.1.3.1 Adhesion to Host’s Cells and Intercellular Matrix................................................................................ 589 51.1.3.2 Evasion of Immune System and Damage of Host’s Tissues.................................................................. 590 51.1.3.3 Resistance to Antimicrobial Drugs........................................................................................................ 591 51.1.4 Microbial Diagnosis.......................................................................................................................................... 591 51.2 Methods............................................................................................................................................................................ 593 51.2.1 Sample Preparation.......................................................................................................................................... 593 51.2.2 Detection Procedures...................................................................................................................................... 593 51.3 Conclusion and Future Perspectives................................................................................................................................. 594 References.................................................................................................................................................................................. 595

51.1  INTRODUCTION Anaerobic bacteria constitute members of the indigenous microbiota in humans and animals. Although these organisms do not usually cause disease, they have the capacity to take advantage of a weakened host’s immune status (e.g., due to injury and immune-suppressing therapy) and can be involved in either mixed or monomicrobial infections. Species of the genus Prevotella form the indigenous microbiota of the oral cavity and body surfaces in humans and animals. The Prevotella genus is a member of the family Bacteroidaceae and phylum Bacteroidetes. However, some species of Prevotella may cause local infections if the equilibrium of the habitat or the host-bacteria relationship is disrupted. In the last two decades, the taxonomy of the genus Prevotella has progressed slowly. The high degree of heterogeneity observed among these indigenous bacteria, and the similarity with other related species, have resulted in a problematic identification. Moreover, several new species have been partially characterized in recent years by using molecular methods, without previous knowledge about their virulence and ecological or epidemiological aspects, to support a significant role in the pathogenesis of infectious diseases or in metabolic traits of this resident microbiota.

51.1.1  Classification, Morphology, and Biology The taxonomy of the gram-negative anaerobes was relatively cumbersome until the 1990s, when the Bacteroides genus was

rearranged, and the genus Prevotella was formed.1 A heterogeneous group of strictly anaerobic bacteria was described as Bacterium melaninogenicum that represented the core of the genus Prevotella and has undergone deep taxonomical rearrangements. This genus includes gram-negative, strictly anaerobic, nonspore-forming, nonmotile, saccharolytic or moderate, and pleomorphic rods. Colonies on blood agar plates vary from 0.5 to 1.0 mm in diameter and are usually circular, entire, convex, shiny, and smooth. They can be gray, light brown, or black-pigmented colonies, and particularly, coccobacilli that fluoresce red are in the pigmented Prevotella spp.–Porphyromonas spp. group. The minimum medium setup includes nonselective, enriched, brucella base sheep blood agar plate supplemented with vitamin K1 and hemin (BAP), and a kanamycin­vancomycin sheep blood agar (KVLB) for selection of Bacteroides and Prevotella spp. KVLB allows growth and rapid pigmentation of most Prevotella spp., but the concentration of vancomycin (7.5 μg/mL) can inhibit most Porphyromonas spp. The hemolytic activity is variable; strains are able to growth from 25°C to 45°C, but the optimal growth is achieved at 37°C. Species of Prevotella are relatively saccharolytic, and most of them use peptides, free amino acids, nitrogen, and carbon as source of energy. In addition, hemin and menadione are the major requirements for growth, and the main fermentation products are acetic and succinic acids, as well as isobutyric, isovaleric, or lactic acids. Historically, oral Bacteroides were separated in two groups, black-pigmented and nonpigmented rods. Those 585

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producing black- or brown-pigmented colonies on blood agar were called B. melaninogenicus; and the nonpigmented colonies, B. oralis. However, the taxonomic importance of the pigment production was overestimated, since the colonial pigmentation was dependent on the composition of the medium. The ability of most clinically relevant species in producing brown-black pigment in presence of hemin and blood is presented in Figure 51.1. Nowadays, the genus Prevotella is composed of 43 species, occupying several ecological niches in resident microbiota from humans and other animals. Figures 51.2 and 51.3 present the most relevant biochemical tests to identify the medically relevant nonpigmented and pigmented Prevotella. However, various new species have been described within genus Prevotella,2–18 and others have been transferred to different genera.19 Therefore, the taxonomic analysis of these bacteria is effervescent. The indole-positive and moderately saccharolytic P. intermedia group harbors P. intermedia, P. falsenii, P.  nigrescens, and P. pallens.5,18 Prevotella intermedia, P. falsenii, and P.  nigrescens are phenotypically identical,2 whereas P.  pallens is lipase-negative and faintly pigment producing.5 Prevotella disiens is similar, but produces neither pigment nor indole. New groups of Prevotella species have been described in recent years, such as P. bergensis, which forms a cluster with P. buccae, P. dentalis, and P. baroniae.20 Both P. enoeca and P. pleuritidis are very similar21; P. copri, P. veroralis, P. shahii, and P. stercorea are genetically closely related, while the recently described P. amnii is closely related to P. bivia.16

P. pallens

P. nigrescens

P. corporis

Species of the genus Prevotella are part of the indigenous microbiota of mucous membranes, especially from the oral cavity,8,9,15,21 and the respiratory, gastrointestinal, and genitourinary tracts of humans12 and other mammals.4,22 Members of the P. melaninogenica group, as well as P. intermedia and P. nigrescens, are among the first anaerobes to colonize the oral cavities of infants.23 In addition, a new species from riceplant residues (P. paludivivens) was recently characterized. Moreover, genus Prevotella represents the most numerous members of the ruminal microbiota (42%–60%), however, typical ruminal species such as P. bryantii, P. ruminicola, and P. brevis represent 2%–4% of the total microbiota, and it is suggested that species of the resident microbiota in animals remain without a correct speciation.4 Ruminal Prevotella species represent a reclassification of strains formerly classified as “Bacteroides ruminicola,” made up of a group of genetically related species, such as P. albensis, P. ruminicola, P. brevis, and P. bryantii, and the hallmark of these four species is the nutritional versatility to use carbohydrates and amino acids.22 The indole-negative and lactose-fermenting P. melaninogenica group includes the phenotypically similar species, such as P. melaninogenica, P. loescheii, and P. denticola.1 Closely related to these species are P. oralis, P. buccae, P. dentalis, P. veroralis, P. bergensis, P. salivae, P. multiformis, P. baroniae, P. oulorum, P. maculosa, P. marshii, P. micans, P. multisaccharivorax, P. nanceiensis, P. shahii, P. paludivivens, and P. timonensis.1,6–8,10,11,14,17,20,21 In addition, oral Prevotella are inhibited by 20% bile and 6.5% NaCl, whereas species belonging to the intestinal microbiota are resistant to both substances. Most of strains grow properly in pH values

P. denticola P. falsenii

P. histicola

Pigmented Prevotella

P. micans P. melaninogenica

P. loescheii

P. intermedia

P. stercorea P. shahii P. salivae P. ruminicola P. pleuritidis P. paludivivens P. oulorum P. oris P. oralis

P. timonensis P. veroralis P. zoogleoformans P. baroniae P. bergensis

Non-pigmented Prevotella

P. buccae P. buccalis

P. copri P. disiens P. enoeca P. multiformis P. heparinolytica P. multisaccharivorax

FIGURE 51.1  Major pigmented and nonpigmented species of genus Prevotella. P. corporis and P. histicola are variable in producing black-pigmented colonies.

587

Prevotella

Fermentation of sucrose –

+ Fermentation of arabinose –

+ Indole – Zoogleal mass in broth –

Zoogleal mass in broth +

P. heparinolytica

–

+ P. zoogleoformans

Fermentation of salicin

+

–

P. zoogleoformans Fermentation of salicin

Fermentation of cellobiose –

+

–

+ P. oralis

+

P. shahii α-Fucosidase β-N-acetyl-glucosaminidase

– P. buccae

+

Fermentation of xylan – P. buccalis

P. oris

+ P. veroralis

FIGURE 51.2  Identification of clinically relevant nonpigmented Prevotella.

varying from 5.7 to 6.7. This genus harbors a heterogeneous group of species, and the different habitats of Prevotella species are presented in Table 51.1. Its members are associated with opportunistic anaerobic infections in oral cavity and in other body sites, and they have been implicated as causative agents of several infectious processes. The infections most commonly associated with genus Prevotella are listed in Table 51.2.

51.1.2  Clinical Features In general, infections caused by anaerobes are endogenous, and these organisms are particularly important in the ensuing phase, characterized by suppuration and abscess formation. The infections involving Prevotella species are similar to other infectious processes associated with gram-negative anaerobes. Usually, the infections involving P. intermedia and P. melaninogenica display acute clinical symptoms, especially in endodontic and periapical infections.37 Most of infectious processes associated with Prevotella spp. are observed in the oral cavity, especially in odontogenic infections. These infections are usually acute and suppurative, showing predominance of Peptostreptococcus, Fusobacterium, and both black-pigmented Prevotella and Porphyromonas species.27 Moreover, the frequency of some

uncultured Prevotella strains from head and neck infections suggests that these anaerobes can play a key role in the pathogenesis of these processes. The presence and intensity of pain, especially in dental pulpitis, is associated with the presence of P. intermedia and other gram-negative anaerobes. The alogenic metabolites from anaerobic gram-negative bacteria in deep caries may partially explain why P. intermedia and the amount of lipopolysaccharide (LPS) in caries are positively related to heat sensitivity or pain. In addition, LPS activates the Hageman factor, leading to bradykinin production, a potent pain inducer. A remarkable characteristic of Prevotella infections is the presence of malodor, and this feature seems to be associated with the production of volatile sulfur compounds by the aminoacid fermentation process of these anaerobes. Among the alogenic molecules produced by the metabolism, ammonia is the most potent pain inducer, followed by urea and indole. Many Prevotella species produce indole from tryptophan. Moreover, Prevotella species, especially P. intermedia, have a significant ability to induce abscess formation, and this characteristic seems to strengthen the virulence of other anaerobes, such as Parvimonas micra.38 Thus, the presence of pus is a hallmark of the infectious diseases in which large numbers of these gram-negative anaerobes are found.28,39

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Molecular Detection of Human Bacterial Pathogens

TABLE 51.1 Ecology of Genus Prevotella

Fermentation of glucose

–

+

Prevotella Species

–

P. intermedia / P. nigrescens/P. falsenii

Fermentation of lactose

–

+

P. corporis Esculin hydrolysis

+ Fermentation of cellobiose

– P. denticola

+ P. loescheii

Rumen Unknowna Oral cavity Unknownb Human mucosa, especially oral cavity and genital tract P. brevis Rumen P. bryantii Rumen P. buccae Oral cavity P. buccalis Oral cavity P. copri Intestines P. corporis Human mucosa P. dentalis Human mucosa, especially oral cavity P. denticola Human mucosa, especially oral cavity P. disiens Human mucosa, especially oral cavity and genital tract P. enoeca Oral cavity P. falsenii Oral cavity from nonhuman primates P. heparinolytica Oral cavity P. histicola Oral cavity P. intermedia Human mucosa, especially oral cavity P. loescheii Human mucosa, especially oral cavity P. maculosa Oral cavity P. marshii Oral cavity P. massiliensis Unknownc P. melaninogenica Human mucosa, especially oral cavity P. micans Oral cavity P. multiformis Oral cavity P. multisaccharivorax Oral cavity P. nanceiensis Human mucosa P. nigrescens Human mucosa, especially oral cavity P. oralis Human mucosa, especially oral cavity P. oris Human mucosa, especially oral cavity P. oulorum Oral cavity P. pallens Oral cavity P. paludivivens Plant residues in the soil P. pleuritidis Unknownd P. ruminicola Rumen Prevotella salivae Oral cavity Prevotella shahii Oral cavity P. stercorea Intestines P. tannerae Oral cavity P. timonensis Unknowne P. veroralis Human mucosa, especially oral cavity P. zoogleoformans Oral cavity P. albensis P. amnii P. baroniae P. bergensis P. bivia

Indole

+

Major Habitat

– Fermentation of cellobiose

– P. melaninogenica

+ P. loescheii

FIGURE 51.3  Identification of clinically relevant pigmented Prevotella.

In oral infections, especially periodontal infections, such as adult and advanced periodontitis, P. intermedia and P. nigrescens play an important role as pathogens and in early periodontal disease without deep periodontal pocket, where Porphyromonas gingivalis, an important periodontopathic bacteria is absent, since Prevotella species are more oxygen tolerant and can provide environmental conditions suitable to supporting other anaerobes. It is well established that periodontal diseases are associated with a complex microbiota, and several bacterial species associated with periodontal diseases have been isolated and characterized,32,40 mainly gram-negative anaerobic microorganisms, including P. gingivalis, Tannerella forsythia, Treponema denticola, P. intermedia, and Fusobacterium nucleatum.41 Several virulence factors in P. intermedia, such as enzyme production, including aminopeptidase-, chymotrypsin-, elastase-, and trypsine-like dipetidylpeptidase and alkaline phosphatase activities have been reported, and these can be harmful to the host.42–46 Oral species of P. intermedia show the ability to adhere and to invade epithelial cells. Leung et al.47 have suggested that the difference in adherence properties might be due to different types of fimbriae present in P. intermedia strains. Dorn et al.48 demonstrated that P.  intermedia invade KB cells and can be internalized within vacuoles. However, the invasion process can be

a b c d e

Isolated from human amniotic fluid. Isolated from skin and soft tissues infections in humans. Isolated from bacteremias in humans. Isolated from human suppurative pleuritis. Isolated from human breast abscess.

Reference  4 16  7 20 24  4  4  1  1 12  1 19 19 25  3 18  1 16  1 19 21  7 26  1 17  8  9 10  2  1  1  1  5 14 13  4  6  6 12  3 11 19  3

589

Prevotella

inhibited by cycloheximide (an inhibitor of protein synthesis in mammalian cells), by cytochalasin D (an inhibitor of actin polymerization), and by monodansylcadaverine (an inhibitor of receptor-mediated endocytosis). In addition, the type C fimbriae of P. intermedia might be a key factor for the invasion since anti-type-C fimbriae antibodies effectively inhibit the P. intermedia invasion.47 Studies in vivo have shown that P. intermedia as well as other periodontal pathogens can be detected in atherosclerotic arteries.49–51

TABLE 51.2 Most Common Infectious Processes Associated with Prevotella Species Infectious Disease or Clinical Condition

Microorganism P. intermedia, P. nigrescens Prevotella spp. Prevotella spp. Prevotella spp. Prevotella spp. Prevotella spp. Prevotella spp. Prevotella spp. Pigmented Prevotella Prevotella spp., particularly P. intermedia and P. nigrescens P. bivia

P. intermedia

P. intermedia Prevotella spp.

Reference

Dental abscesses Breast abscesses Skin abscesses Bacteremias and septicemias Amniotic infections Bone infections Pleural infections Bite wound Otitis Endodontic and periapical infections

27,28 11 20 26

Gynecology and obstetric diseases in patients evidencing preterm delivery Periodontal infections: chronic periodontitis, necrotizing gingivitis, pregnancy gingivitis Oral malodor Upper respiratory tract infections

24

Tissue necrosis and delay of wound

Low weight toxic compounds (H2S, ammonia)

Anaerobes implicated in infectious diseases are usually in association with facultative and other anaerobic bacteria, and these infections depend on the adhesion to the host’s cells. After these initial steps, tissue invasion may occur. However, the bacteria in the front of the invasion need to survive against the host’s defenses and, at the same time, acquire nutrients to stay alive, and this process frequently ends with necrosis. In this sense, Prevotella species have a significant range of virulence factors that allow them to adhere to and invade the host’s tissues, destroy cellular structures and matrix, and stimulate a powerful and persistent inflammatory process. Figure 51.4 summarizes most of the virulence mechanisms of Prevotella species involved in human infections.

32–35

51.1.3.1  A  dhesion to Host’s Cells and Intercellular Matrix Bacterial adherence to host’s cells is the initial step in colonization. Besides the adhesion process, the bacterial

36 29

Evasion of immune system Pain induction

51.1.3  Virulence and Pathogenesis

16 25,29 13 29 30 27,28,31

Persistence in tissues

Invasion of host’s cells

Biofilm development

Adhesion to glucosamine or galactosides residues

Amino acid fermentation

Resistance to antimicrobials

Degradation of proteins and immunoglobulins

Tissue necrosis and/or osteolytic lesions

Anti-phagocytic capsule

Genus Prevotella

Proteases and peptidases

Invasion and dissemination

Co-aggregation with microorganisms

Hemolysins and cytolysins

LPS Bradikynin production Pain

Cell damage and inflammation

Induction of inflammation

FIGURE 51.4  Overview of the virulence of Prevotella species.

Tissue damage

Evasion of Immune system

Iron source

Virulence control

Evasion of immune system

Persistence and dissemination

590

coaggregation is extraordinarily important in the development of microbial communities involved in mixed anaerobic infections. Studies on the adhesion of Prevotella are considered for oral species, particularly P. intermedia, P. loescheii, and P. nigrescens. Many Prevotella species possess fimbriae, adhesins, and hemagglutinins.52,53 The adhesins and hemagglutinins of P. loescheii and P. melaninogenica have affinity to galactose-containing carbohydrates residues on the host’s cells; whereas, in P. intermedia and P. nigrescens, the adhesion molecules interact with glucosamine-containing carbohydrates. In addition, the hemagglutinating activity of Prevotella species is not mediated by proteolytic activities, unlike that P. gingivalis. In addition, P. intermedia and P. nigrescens are able to adhere to large varieties of cells and proteins, such as fibroblasts and reconstituted membranes, epithelial cells, and erythrocytes,54 as well as coaggregate with other microorganisms as Actinomyces, Fusobacterium, Porphyromonas, and Streptococcus.55 Prevotella intermedia is able to coaggregate with several other early colonizers, such as Streptococcus salivarius,56 and creating conditions for the establishment of some late colonizers, as P. gingivalis.57 The coaggregation with P. gingivalis may be inhibited by addition of l-arginine and l-lysine. Prevotella loescheii produces a galactoside-specific adhesin that mediates the hemagglutination of neuraminidase-treated human erythrocytes and lactose-inhibitable coaggregations with oral streptococci. The adhesin is a 75-kDa protein that is expressed on the cell surface in a maximum of 400 molecules per cell, and this adhesin is associated with the distal portion of the fimbriae. A second adhesin of 45 kDa that mediates lactose-noninhibitable coaggregation with A. israelii is also expressed by P. loescheii.55 The coaggregation partners of oral Prevotella species appear to be primarily the actinomycetes. Prevotella produces a coaggregation by adhesins and actinomyces by carbohydrate-containing receptors on its cell surface. In addition, a protease synthesized by P. loescheii is able to degrade a 75-kDa adhesin, suggesting that proteases may actually aid in detachment of this bacterium from the coaggregation partner cell, allowing finding a different niche in oral biofilm.55 51.1.3.2  E vasion of Immune System and Damage of Host’s Tissues In order to produce infections, a microorganism must be able to adhere to hosts’ cells and colonize them, as well as produce harmful compounds and toxins to induce damage or be capable of invading tissues. In addition, evasion of immune defenses represents major strategy for microbial dissemination and persistence in the hosts, and several mechanisms may be involved in this phenomenon. Capsule. Capsule is believed to help bacteria avoid phagocytosis and killing by polymorphonuclear leukocytes (PMNs) by preventing the deposition of opsonins on the bacterial surface or by mimicking mammalian cell-surface determinants.58 Encapsulated anaerobes generally induce abscess, but nonencapsulated organisms are not able to induce it, as also described to P. melaninogenica, P. oralis, and P. brevis,

Molecular Detection of Human Bacterial Pathogens

and the ability to produce abscess is not associated with the presence of viable cells. Moreover, some species, such as P. intermedia, P. oralis, and P. melaninogenica, may be profusely capsulated. Low-Weight Metabolic End Products, Lipopoly­ saccharides, Toxins, and Induction of Inflammation. These anaerobes are able to produce volatile sulfur compounds (hydrogen sulfide and methylmercaptans) associated with malodor, which may reduce the cellular proliferation and repair. Volatile sulfur compounds are produced by anaerobic bacteria derived from amino acids desulfuration containing sulfhydryl groups. For instance, hydrogen sulfide is derived from desulfuration of cysteine; and methylmercaptan, from methionine desulfuration. Thus, both P. intermedia and P. nigrescens are potent producers of these compounds.59 Species of Prevotella produce succinate, acetate, and isovalerate with a small amount of isobutyrate in the absence of glucose, but produce a large amount of ammonia, which has a significant immunosuppressive effect and delays the wound repair. These short-chain fatty acids are reported to inhibit the function and/or proliferation of neutrophils, T-lymphocytes, phagocytes, and fibroblasts.60 The presence of glucose represses the intracellular enzymes formation involved in the nitrogen-source metabolism. Thus, in the oral cavity around supragingival biofilm, where saliva and diet residues are available for the microbial metabolism, the fermentation of carbohydrates leads to environmental acidification, and the oxygen can reach the bacteria around the gingival margin. However, P. intermedia and P. nigrescens can survive and even grow under these conditions because they are saccharolytic, acid-tolerant, and oxygen-tolerant61; but in the bottom of the periodontal pockets and in deep internal infections, where carbohydrate supply is limited, the pathogenicity of P. intermedia, and P. nigrescens may be constantly elevated.60 Markers of systemic inflammation, such as the presence of C-reactive protein in plasma are indicative of myocardial infarction or ischemic stroke risk. Although several factors may increase plasmatic concentrations of C-reactive protein, this presence of P. intermedia and other anaerobes in subgingival samples was positively associated with elevated levels of C-reactive protein.62 Since one of the hallmarks of periodontal disease is inflammation, mainly in chronic periodontal inflammation, it is possible that periodontal pathogens may be involved with the increase in C-reactive levels. Lipopolysaccharide is a major component of the outer membrane of gram-negative bacteria, including Prevotella species. It has the ability to trigger a number of host cells, especially mononuclear phagocytes, to produce a wide variety of immunologically active mediators, including TNF-α, interleukins, and nitric oxide.63 Moreover, LPS produced by P. intermedia may participate in the periodontal destruction and alveolar bone loss through the osteoclastogenesis stimulation,64 as well as reduce bone formation. However, endotoxins of Prevotella spp. are less potent than endotoxins extracted from Enterobacteriaceae and are less likely to produce classic manifestations of toxic damage.58

591

Prevotella

Tissue Invasion. The ability to invade host tissues may be an important virulence factor in bacteria. P. intermedia can invade oral epithelial cells, but the type C fimbriae and a cytoskeleton rearrangement are required for invasion.48 Moreover, P. bivia is also capable of adhering to and invading human cervix epithelial cells, and of inducing proinflammatory interleukins (IL-6 and IL-8), but this phenomenon is not related to the presence of fimbriae.65 In addition, DNA of P. intermedia may be present in atherosclerotic plaques, and this anaerobe may play a role in development and progression of atherosclerosis, leading to a possible coronary vascular disease or other clinical sequel, suggesting a possible adhesion and invasion to the coronary cells.66 Some evidence suggests that the absence of hemin may limit the ability of P. intermedia to invade dentinal tubules.67 However, the mechanism by which the iron interferes with invasion is not yet known. In theory, bacterial attachment to dentine and tubule invasion requires of adhesins, and it is known that stressed cells produce a different protein profile, particularly, to the cell survival. Hemolysis. Another virulence factor is a cytolytic molecule with hemolytic activity that seems to contribute to the pathogenicity of these microorganisms by favoring the iron acquisition that is essential to their metabolism, to survival, and to act upon other host cells.68 Prevotella intermedia lack the ability to synthesize heme, which is an important growth factor.69 P. intermedia can use hemoglobin as a sole source of iron, and inorganic iron and iron-binding proteins, such as transferrin and lactoferrin, could not support the bacterial growth, suggesting that hemoglobin might be the main iron source. However, the mechanisms to the iron assimilation from hemoglobin have not been yet elucidated. The whole cells of P. intermedia have a weak hemolytic activity on human erythrocytes, but when these microorganisms are lyzed and hemolysin-free and are treated by trypsinlike proteases, the lytic activity is significantly increased. In contrast, no increase in the lytic activity of P. nigrescens was observed after rupturing the bacteria, although proteolytically treated hemolysin produce a very high intensity of hemolysis, higher than P. intermedia hemolysin.68 Proteolytic Activities. Prevotella species produce a wide variety of enzymes associated with evasion of immunological responses or tissue damage. In addition, cysteine protease (linked to hemoglobin degradation) and other cysteine and serine proteases have been described.70,71 All these proteases have been identified as secreted virulence factors producing nutrient generation, evasion of the immune response through the immunoglobulin inactivation, and by releasing bacterial proteins from cell surface. Immunoglobulin A1 protease, secreted by most oral Prevotella species, particularly P. melaninogenica, P. intermedia, and P. nigrescens, specifically cleaves IgA1 at the same peptide bond in the hinge region, and this phenomenon occurs in vivo, especially in periodontitis patients. However, human serum from these patients present high titles of neutralizing antibodies able to inhibit partially the proteolytic activity of IgA1 proteases produced by Prevotella species,

such as P. buccae, P. oris, P. loescheii, P. buccalis, and P. melaninogenica.72 These enzymes may improve the pathogen dissemination, as well as provide amino acids used in the catabolic reactions of these anaerobes. Some evidences suggest that P. loescheii, P. melaninogenica, P. intermedia, and P. oralis recovered from dentoalveolar abscesses are more proteolytic than those obtained from healthy periodontal sites. In addition, P. intermedia and P. nigrescens are the most proteolytic among the oral Prevotella species.73 Moreover, serine proteases and exopeptidases produced by P. loescheii have been described recently, which help trypsin- and collagenase-like protease in the type I collagen degradation, but they are strongly inhibited by Cd2+, Zn2+, Hg2+, Co2+, and Cu2+. Whole-cell extracts of P. nigrescens activate the host’s metalloproteinases responsible for the matrix proteins’ depolymerization and the dissemination of these anaerobes in connective tissues.74,75 Most of studies on proteolytic activity have been performed with oral Prevotella and ruminal species, particularly, P. ruminicola, P. brevis, P. albensis, and P. bryantii, that have also evidenced strong proteolytic abilities, mainly on casein, and several proteases such as serine, cysteine, and low molecular weight metalloproteases were detected.76 51.1.3.3  Resistance to Antimicrobial Drugs Bacteria of the genus Prevotella are often resistant to antimicrobials, such as tetracycline,77 clarithromycin, quinolones,78 β-lactam antibiotics,28,39,79 and clindamycin.77 The β-lactam resistant Prevotella strains are β-lactamase producers, especially P. bivia, P. intermedia, P. nigrescens, P. melaninogenica, P. loescheii, P. oralis, P. buccae, and P. oris.79,80 β-lactamases enzymes produced by these anaerobes display properties of the class A-group 2e β-lactamases, which hydrolyze most penicillins and broad-spectrum cephalosporins but are inactive against penicillin plus clavulanic acid and carbapenems.79 Moreover, it has been evidenced that oxygen exposition may affect the susceptibility of P. intermedia to antimicrobial drugs, probably due to upregulation of protective enzymes by oxidative stress.61

51.1.4  Microbial Diagnosis The purpose of the microbial identification is to match an isolate with a previously recognized taxonomic group, using a number of phenotypic or genetic characters that can be evaluated. The identification systems must be reliable, convenient, rapid, easily performed, flexible, and cheaper. Conventional Techniques. Usually, the conventional techniques for the diagnosis of anaerobic infections depend, at least to some extent, on culture. However, variations of environmental parameters, including redox potential, osmolarity, pH, temperature, and nutrients can interfere with detection of a pathogen. In microbiological diagnostics, the bacterial culture has been the gold standard for many years, although limitations with respect to detecting nonviable bacteria and the inability of some species to grow on selective

592

media, as well as the high cost, are known. Despite their effectiveness and sensitivity, these procedures are extremely labor intensive and time consuming. Moreover, classification and identification based on phenotypic traits do not always provide clear-cut results and are sometimes unreliable. The definitive identification of anaerobes is based on the use of biochemical tests in combination with gas–liquid chromatography, but this method is rather labor intensive and expensive. Thus, the development of automated methods for anaerobes identification has received interest. Some systems employ computerized high-resolution gas–liquid chromatography to determine the fatty acid composition from bacteria, which is then compared with a database. However, the performance of these systems is precarious for Prevotella species.81 Methods used to identify Prevotella strains includes colony morphology, pigmentation, Gram staining, carbohydrate fermentation, esculin hydrolysis, gelatin liquefaction, nitrate reduction, and indole production in PRAS media, and the detection of preformed enzymes by using RapID ANA II, RapiID 32, and API ZYM kits. Pure cultures must be stored in 20% skim milk at –80°C or liquid nitrogen. Using filter-paper spot tests for indole production, sialidase, α-glucosidase, β-glucosidase, α-fucosidase, and trypsin-like enzyme activities, the presumptive identification of oral Prevotella may be achieved. The bacterial pigmentation is largely media dependent, and the pigment production is used for distinguishing colonies in early stages of identification. The screening using rapid tests is enough for identifying the most common oral Prevotella species, such as P. intermedia, P. nigrescens, and P. melaninogenica. In addition, the indole test would increase the accuracy of a simple identification scheme by recognizing occasional β-galactosidase (MUG)-positive P. intermedia/P. nigrescens strains, otherwise misidentified as belonging to the P. melaninogenica group. Glucose, sucrose, and lactose fermentation, pigment, indole, and lipase production, as well as α-fucosidase and N-acetyl-β-glucosaminidase assays are the core test for identification of P. intermedia/P. nigrescens group, allowing the partial identification of P. pallens, P. intermedia, and P. nigrescens, P. corporis, and P. disiens.5 Other phenotypical tests useful in the identification of Prevotella spp. include gelatin and esculin hydrolysis.6 On the other hand, ambiguous results are often obtained due to the emergence of phenotypically variable strains.5,8 Moreover, the speciation of some species, such as P. intermedia, P. nigrescens, and P. tannerae, needs additional molecular tests, particularly, 16S rDNA-based polymerase chain reaction (PCR), since some black-pigmented Prevotella have been previously identified as P. intermedia by using rapid biochemical identification kits, and then as P. nigrescens by using SDS-PAGE. Thus these species were identified as P. tannerae when PCR and restriction enzyme analysis were performed. Molecular Techniques. Molecular detection methodologies are powerful tools used for microbial identification, to

Molecular Detection of Human Bacterial Pathogens

study host-parasite relationships, and to clarify taxonomic positions of targeted microorganisms. Most of molecular techniques used in the microbial diagnosis are based on PCR, which can unequivocally detect phenotypically variable isolates and can be considered as a more specific and reliable method for bacterial identification; otherwise, it is a rapid and relatively inexpensive identification method. Nowadays, the 16S rRNA gene PCR is considered as an accurate and reproducible method for separation among P. nigrescens and P. intermedia, as well as to identify the Prevotella species directly from clinical samples.82–84 Moreover, other genes have been targeted with speciesspecific PCR, such as genes phoC (P. intermedia acid phosphatase), phyA (P. melaninogenica hemolysin), and plaA (P. loescheii adhesin precursor).85 The ribosomal RNA genes and elongation factor G have provided valuable phylogenetic information for anaerobes and other microorganisms. The 16S rRNA gene has been widely used for phylogenic studies, and has produced phylogeny comparable to the 23S rRNA genes. The phylogenetic analysis of Prevotella species have indicated some variations in the 16S rDNA sequences within the genus, and it is enough to make the 16S rRNA gene analysis a good tool for the species-specific PCR identification, both from cultures and directly from clinical samples.82,86 In addition, the 16S rDNA sequences have been the target for simultaneous detection of different species by multiplex PCR, including P. nigrescens, although other genes have been detected by using multiplex PCR in P. intermedia, P. melaninogenica, and P. loescheii.85 Other molecular methods have been used to separate clinically relevant Prevotella species, particularly, P. nigrescens from P. intermedia, such as multilocus enzyme electrophoreses,87 restriction enzyme analysis of total DNA, ribotyping,88 SDS-PAGE protein electrophoresis,87 16S rRNA gene PCR-restriction fragment length polymorphism,89 and 16S rRNA gene cloning and sequencing.36,39 The real-time PCR assays for Prevotella species allow an easy, rapid, and quantitative detection of these microorganisms directly from clinical samples. The TaqMan system is the most frequently used and confers high sensitivity and specificity to DNA amplification. Primers and probes sets for Bacteroides-Prevotella group,90 P. ruminicola group,22 P. intermedia, and P. nigrescens,84 P. melaninogenica, and P. loescheii,91 P. melaninogenica, and Prevotella sp.22,83 have been described. In addition, protocols for quantitative detection of genes resistant to antimicrobials are also available.79 Real-time PCR combines traditional end-point-detection PCR with fluorescent detection technologies to record the accumulation of amplicons in real time during each cycle of the PCR amplification. The detection of amplicons during the early exponential phase of the PCR enables the quantification of gene (or transcript) numbers when these are proportional to the starting template concentration. This method has been shown to be a robust, highly reproducible, and sensitive method of quantifying functional genes in many conditions. The success of the application

593

Prevotella

of this methodology is based on the specificity of real-time PCR assay, which is determined by the design of the primers (or internal probes). Ribosomal RNA or ribosomal 16S (16S rRNA) (specific prokaryote rRNA) is the most widely used macromolecule in phylogenetic and taxonomic studies. 16S rRNA analysis has been widely used to establish phylogenetic relationships in the world of prokaryotes, exerting a profound impact on our view of evolution and, as a result, on the classification and identification of bacteria. One of the first techniques used in molecular biology to detect target microorganisms in clinical samples was the nucleic acid probe. A probe is formed by a short DNA fragment, containing only a few nucleotides, complementary to the DNA belonging to the target, and should be labeled with a molecule that is easily detected once hybridization has occurred. The main problem with the technique is the risk of cross-reaction. The checkerboard DNA–DNA hybridization technique allows the enumeration of large numbers of species in very large numbers of samples, and 16S rRNA gene clone library analysis can be used to survey the diversity of human oral microbiota and the existence of as-yetto-be-cultured organisms that are presumed pathogens. In addition, terminal restriction fragment length polymorphism (T-RFLP) analysis has been also applied for assessment of diversity of human oral microbiota.

51.2  METHODS 51.2.1  Sample Preparation The clinical samples commonly used to detect species of Prevotella are pus,31,39,80 necrotic tissues,80 biopsied tissues,25 vaginal discharge,24 and dental biofilm.85 Bone infection clinical samples, especially of the skull bones, must avoid contact with external contaminants, and only five types of clinical specimens are acceptable for microbial diagnosis: bone biopsy, bone sequestra, bone marrow, granulation tissue, and aspirated pus. In order to avoid the artificial introduction of oral microorganisms into specimens, it now required that clinical specimens must be taken during surgical procedures, avoiding the contact with soft tissues, the oral environment, and sinus tracts. Whenever possible, bone sequestra or granulation tissue samples must also be obtained for the histopathological examination. The best strategy for collecting pus is needle aspiration and transportation inside a prereduced medium. The use of transport medium is unnecessary for short periods of time.25,39 In vaginal infections, the use of cotton wool swabs is suitable when vaginal discharge is present.24 For isolation of Prevotella species from rumen liquids and contents, samples22 must be transported in an environment containing only CO2 to avoid contact with oxygen or in prereduced transport media. The most often used prereduced transport media for transportation of specimens containing anaerobic bacteria, including samples with Prevotella species, are Ringer-PRAS, RTF,83,92 and VMGA III.37,92 In addition, since preparation, storage, and handling of VMGA III medium are reliable, easily performed, and support the survival of anaerobic bacteria

for 1 or 2 days, this medium is widely used as a standard in clinical laboratories. In oral samples, it is important to note when antiseptics were used to avoid sample collection if required, because the antimicrobial action can interfere with the bacterial recovery. Thus, use of sterile 5% sodium thiosulfate is recommended to eliminate residual antimicrobials.

51.2.2  Detection Procedures The use of conventional techniques for the analysis of infections involving Prevotella species has shown limitations due to their susceptibility of the molecular oxygen. Strictly anaerobic bacteria are not equally susceptible to oxygen, and even different strains may display different levels of sensibility, which is reduced by continuous subcultures and by the production of protective enzymes acting against the toxic products derived from oxygen reduction. The ability to adapt to the oxygen changes is a remarkable physiologic trait of pathogenic Prevotella.61 However, the prime isolation of these microorganisms is Eh-negative dependent, since fresh isolates are more susceptible to the oxygen’s producing a reduction of 90%–99% of the microbial number after 15 h of exposition to molecular O2. Traditionally, culture for anaerobes is a time-consuming and expensive procedure. Most clinical and research laboratories use the so-called strictly anaerobic techniques or conventional anaerobic techniques for isolation of Prevotella species. These techniques differ from those considered “conventional” because the specimen is never exposed to air during the entire process of preparation and culturing, or at least the contact with molecular oxygen is minimized by using PRAS transport and dilution solutions, and the media used are also prereduced. All pathogenic Prevotella species grow on nutritionally rich culture media, such as brucella agar, Columbia agar, brain heart infusion agar, fastidious anaerobe agar, or even trypticase soy agar, supplemented with hemin (0.05 mg/dL), menadione (0.01 mg/dL), and yeast extract (0.1%–5%). However, since most of Prevotella species are members of human and animal resident microbiota, their isolation requires the use of selective culture media, such as KVLB agar, in which kanamycin and vancomycin exert a selective role, especially for oral Prevotella. The association of 5% sheep or horse blood with 0.05 mg/dL hemin, 0.01 mg/dL vitamin K1, 0.001% w/v nalidixic acid, and 2.5 mg/L vancomycin using WilkinsChalgren agar is also adequate for isolation of these anaerobes. Although, some Prevotella species are able to produce visible growth on agar plates after 48 h, the incubation must be performed during 7–15 days, to allow the development of the most fastidious species and the brown-pigment characteristic observed in the most virulent species. The detection procedures of the molecular methods do not have specific peculiarities in relation to facultative anaerobes and other anaerobes. However, DNA extraction by heat (96°C, for 10 min) has been used with Prevotella spp.,79 and in some circumstances DNA extraction and amplification may be affected by the presence of cellular debris and iron

594

Molecular Detection of Human Bacterial Pathogens

because they may affect the DNA amplification by chelating magnesium or interfering with DNA polymerase. To minimize this problem, the DNA extraction must be performed by using other methods, providing an additional clearance, such as phenol-chloroform or commercial kits. In addition, due to the nature of PCR-based protocols with high levels of sensitivity, the main shortcomings in applying PCR assays include false-positive results due to background DNA contaminations. These concerns are particularly relevant in the microbial diagnosis of members of the human and animal resident microbiota. In addition, the presence of one microorganism is not necessarily an indication that it plays a pathogenic role in the infection development, and this is true in mixed anaerobic infections, where different microorganisms share the same habitat. Bacterial anaerobic culture is based on living cells. This technique enables us to search for all the microorganisms present in a nonspecific way. Since it is not directed toward precise pathogenic targets, it remains the most objective technique (gold standard), but requires at least 103 pathogens for detection. Thus, many unexpected pathogen colonies appear during culture, whereas routine probes cannot detect them. Also, the cultures have the important advantage of allowing an antibiotic sensitivity. If we consider the interindividual or the intraindividual variations in the susceptibility of clinical isolates to antibiotics, in particular for P. intermedia, there is a good reason for to perform the conventional bacterial culture and antibiogram from each patient. On the other hand, the detection of pathogens by the real-time PCR is obtained even when they are at low levels, showing high sensitivity, and because the technique detects the DNA of both living and dead bacteria. Most of studies on detection of Prevotella species were carried out using oral specimens, particularly from periodontal and endodontic/periapical infections, where P. intermedia, P. nigrescens, P. melaninogenica, and P. loescheii are common. Based on the four Prevotella species-specific genes, Yoshida et al.85 described one multiplex PCR focusing on P. intermedia, P. nigrescens, P. melaninogenica, and P. loescheii from oral cavity (Table 51.3).

Procedure.  The oligonucleotide primers and their DNA sequences are listed in Table 51.3, and the specificity of the primer pairs is confirmed by sequencing. Multiplex PCR for black-pigmented oral Prevotella may be performed as follows: 95°C for 15 min, followed by 35 cycles of 94°C for 30 s, 54°C for 90 s, and 72°C for 90 s. A 50-µL PCR mixture for multiplex PCR consists of 25 µL 2× Qiagen Multiplex PCR Master Mix (Qiagen), 2 µM of each primer, and 1 µL of template DNA, and 1–10 ng of genomic DNA as template. After completion of PCR cycles, add 4 μL of 10× DNA loading buffer into each tube; load 20 μL of PCR product on 1.0% 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 transilluminator and photograph.

51.3  C  ONCLUSION AND FUTURE PERSPECTIVES Almost all available knowledge about the physiology of these anaerobes is based on data obtained from oral Prevotella species, particularly P. intermedia, but this microbial genus is extremely heterogeneous, not only from the phenotypical level, but also mainly at the molecular level. Thus, it is entirely possible that new interactions between the newly described species and theirs hosts may be characterized in the coming years. These ecological interactions may explain the mechanisms involved in the pathogenicity of these anaerobes and may collaborate in the development of more ecological therapeutic approaches in the treatment endogenous infections associated with these microorganisms, and may also collaborate for a better understanding of the role played by these microorganisms as part of the resident microbiota of the oral cavity and other mucous membranes. The characterization of Prevotella paludivivens, a hemicellulose-decomposing bacterium isolated from plant residue and rice roots in irrigated rice-field soil, opens the possibility of the participation of this strictly anaerobic genus in complex microbial communities not related to infections. The ability of these environmental microorganisms also represents an

TABLE 51.3 Multiplex PCR for Black-Pigmented Prevotella from Oral Cavity Prevotella Species

Sequence of Oligonucleotide Primers

Gene

P. intermedia

5′-GAC TTT TGC ACA GAA TGC A-3′ 5′-CTT GGC AAC CTT GCC TTC-3′ 5′-TGC CAA CTC CCG ATT TC-3′ 5′-TAC ACC AAG GTT TTC CCC-3′ 5′-CGT CAT GAA GGA GAT TGG-3′ 5′-ATA GAA CCG TCA ACG CTC-3′ 5′-TTT CAT TGA CGG CAT CCG-3′ 5′-CAC GTC TCT GTG GGC AG-3′

phoC

718

plaA

637

phyA

389

16S rRNA

825

P. loescheii P. melaninogenica P. nigrescens

Source: Yoshida, A. et al., Oral Microbiol. Immunol., 20, 43, 2005.

Amplicon Size (bp)

595

Prevotella

exceptional opportunity to study the similarities and differences between species frequently isolated from infectious processes and those recovered from symbiotic communities in the external environment. In addition, approximately three-quarters of all species of the genus Prevotella were identified and characterized during studies with clinical specimens from the human oral cavity and from infections, and almost nothing is known about the anaerobic microbiota from other animal species, even the nonhuman primates that are often used as animal models in studies involving medicine, veterinary medicine, and dentistry. Thus, it is likely that in the future, the number of species of this genus from animal origin will increase significantly, just as the knowledge available on the potential virulence of these microorganisms has. Nowadays, several markers of resistance to antibiotics characteristic of other anaerobic species has been identified in species of the genus Prevotella, but the distribution of major genes associated with resistance to antimicrobials, especially those associated with resistance to β-lactams, tetracyclins, macrolides, and nitroimidazoles among species of the genus Prevotella, particularly P. intermedia and P. nigrescens, is not known, and the possibility of vertical and horizontal transmission of these markers to the other microorganisms inside the dental biofilm and intestinal microbiota remains unexplored. Due to the development of molecular methods, it has become possible to evaluate the factors capable of controlling the expression of genes related to primary and/or secondary metabolism in opportunistic pathogens, enabling a greater understanding of how environmental factors or hosts’ factors may exacerbate or depress the microbial virulence. This aspect seems particularly relevant to the oral species of the genus Prevotella, which are associated with gingival changes modified by hormonal changes. It is possible that this knowledge may enable the development of preventive strategies for numerous diseases associated with hormonal changes.





REFERENCES







1. Shah, H.N., and Collins, D.M., Prevotella, a new genus to include Bacteroides melaninogenicus and related species formerly classified in the genus Bacteroides, Int. J. Syst. Bacteriol., 40, 205, 1990. 2. Shah, H.N., and Gharbia, S.E., Biochemical and chemical studies on strains designated Prevotella intermedia and proposal of a new pigmented species, Prevotella nigrescens sp. nov., Int. J. Syst. Bacteriol., 42, 542, 1992. 3. Moore, L.V., Johnson, J.L., and Moore, W.E., Descriptions of Prevotella tannerae sp. nov. and Prevotella enoeca sp. nov. from the human gingival crevice and emendation of the description of Prevotella zoogleoformans, Int. J. Syst. Bacteriol., 44, 599, 1994. 4. Avgustin, G., Wallace, R.J., and Flint, H.J., Phenotypic diversity among ruminal isolates of Prevotella ruminicola: Proposal of Prevotella brevis sp. nov., Prevotella bryantii sp. nov., and Prevotella albensis sp. nov. and redefinition of Prevotella ruminicola, Int. J. Syst. Bacteriol., 47, 284, 1997.





5. Könönen, E. et al., Phylogenetic characterization and proposal of a new pigmented species to the genus Prevotella: Prevotella pallens sp. nov., Int. J. Syst. Bacteriol., 48, 47, 1998. 6. Sakamoto, M. et al., Prevotella shahii sp. nov. and Prevotella salivae sp. nov., isolated from the human oral cavity, Int. J. Syst. Evol. Microbiol., 54, 877, 2004. 7. Downes, J. et al., Prevotella marshii sp. nov. and Prevotella baroniae sp. nov. isolated from the human oral cavity, Int. J. Syst. Evol. Microbiol., 55, 1551, 2005. 8. Sakamoto, M. et al., Prevotella multiformis sp. nov., isolated from human subgingival plaque, Int. J. Syst. Evol. Microbiol., 55, 815, 2005. 9. Sakamoto, M. et al., Prevotella multisaccharivorax sp. nov., isolated from human subgingival plaque, Int. J. Syst. Evol. Microbiol., 55, 1839, 2005. 10. Alauzet, C. et al., Prevotella nanceiensis sp. nov., isolated from human clinical samples, Int. Syst. Evol. Microbiol., 57, 2216, 2007. 11. Glazunova, O. et al., Prevotella timonensis sp. nov., isolated from a human breast abscess, Int. J. Syst. Evol. Microbiol., 57, 883, 2007. 12. Hayashi, H. et al., Prevotella copri sp. nov. and Prevotella stercorea sp. nov., isolated from human faeces, Int. Syst. Evol. Microbiol., 57, 941, 2007. 13. Sakamoto, M. et al., Prevotella pleuritidis sp. nov. isolated from pleural fluid, Int. Syst. Evol. Microbiol., 57, 1725, 2007. 14. Ueki, A. et al., Prevotella paludivivens sp. nov., a novel strictly anaerobic, gram-negative, hemicellulose-decomposing bacterium isolated from plant residue and rice roots in irrigated rice-field soil, Int. J. Syst. Evol. Microbiol., 57, 1803, 2007. 15. Downes, J. et al., Prevotella histicola sp., nov., isolated from the human oral cavity, Int. J. Syst. Evol. Microbiol., 58, 1788, 2008. 16. Lawson, P.A., Moore, E., and Falsen, E., Prevotella amnii sp. nov., isolated from human amniotic fluid, Int. J. Syst. Evol. Microbiol., 58, 89, 2008. 17. Downes, J. et al., Prevotella micans sp. nov., isolated from the human oral cavity, Int. J. Syst. Evol. Microbiol., 59, 771, 2009. 18. Sakamoto, M. et al., Prevotella falsenii sp. nov., a Prevotella intermedia like organism isolated from monkey dental plaque, Int. J. Syst. Evol. Microbiol., 59, 319, 2009. 19. Willems, A., and Collins, M.D., 16S rRNA gene similarities indicate that Hallella seregens (Moore and Moore) and Mitsuokella dentalis (Haapasalo et al.) are genealogically highly related and are members of the genus Prevotella: Emended description of the genus Prevotella (Shah and Collins) and description of Prevotella dentalis comb. nov., Int. J. Syst. Bacteriol., 45, 832, 1995. 20. Downes, J. et al., Prevotella bergensis sp. nov., isolated from human infections, Int. J. Syst. Evol. Microbiol., 56, 609, 2006. 21. Downes, J. et al., Prevotella maculosa sp. nov., isolated from the human oral cavity, Int. J. Syst. Evol. Microbiol., 57, 2936, 2007. 22. Stevenson, D.M., and Weimer, P.J., Dominance of Prevotella and low abundance of classical bacterial species in the bovine rumen revealed by relative quantification real-time PCR, Appl. Microbiol. Biotechnol., 75, 165, 2007. 23. Kobayashi, N. et al., Colonization pattern of periodontal bacteria in Japanese children and their mothers, J. Periodontol. Res., 43, 156, 2008. 24. Mikamo, H. et al., Bacterial flora in the uterine endometrial cavity of healthy subjects on the first and fifth days of the puerperium, J. Infect. Chemother., 4, 12, 1998.

596 25. Fukuoka, M. et al., Pyogenic vertebral osteomyelitis caused by Prevotella intermedia, Infect. Chemother., 8, 182, 2002. 26. Berger, P. et al., Prevotella massiliensis sp. nov. isolated from human blood, Res. Microbiol., 156, 967, 2005. 27. Gomes, B.P.F.A. et al., Microbiological examination of infected dental root canals, Oral Microbiol. Immunol., 19, 71, 2004. 28. Robertson, D., and Smith, A.J., The microbiology of the acute dental abscess, J. Med. Microbiol., 58, 155, 2009. 29. Brook, I., Microbiology and management of joint and bone infections due to anaerobic bacteria, J. Orthop. Sci., 13, 160, 2008. 30. Brook, I., The role of anaerobic bacteria in chronic suppurative otitis media in children: Implications for medical therapy, Anaerobe, 14, 297, 2008. 31. Al-Nawas, B., and Maeurer, M., Severe versus local odontogenic bacterial infections: Comparison of microbial isolates, Eur. Surg. Res., 40, 220, 2008. 32. Moore, W.E.C., and Moore, L.V.H., The bacteria of periodontal diseases, Periodontology, 5, 66, 1994. 33. Botero, J.E. et al., Occurrence of periodontopathic and superinfecting bacteria in chronic and aggressive periodontitis subjects in a Colombian population, J. Periodontol., 78, 696, 2007. 34. Lafaurie, G.I. et al., Demographic, clinical and microbial aspects of chronic and aggressive periodontitis in Colombia: A multicenter study, J. Periodontol., 78, 629, 2007. 35. Herrera, D. et al., Subgingival microbial profiles in chronic periodontitis patients from Chile, Colombia and Spain, J. Clin. Periodontol., 35, 106, 2008. 36. Haraszthy, V. et al., Identification of oral bacterial species associated with halitosis, J. Am. Dent. Assoc., 138, 1113, 2007. 37. Cogulu, D. et al., PCR-based identification of selected pathogens associated with endodontic infections in deciduous and permanent teeth, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endodontol., 106, 443, 2008. 38. Araki, H. et al., The microbial synergy of Peptostreptococcus micros and Prevotella intermedia in a murine abscess model, Oral Microbiol. Immunol., 19, 177, 2004. 39. Riggio, M.P. et al., Identification of bacteria associated with spreading odontogenic infections by 16S rRNA gene sequencing, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endodontol., 103, 610, 2007. 40. Socransky, S.S., and Haffajee, A.D., The bacterial etiology of destructive periodontal disease: Current concepts, J. Periodontol., 63, 322, 1992. 41. Darby, I., and Curtis, M., Microbiology of periodontal disease in children and young adults, Periodontology, 26, 33, 2001. 42. Slots, J., Enzymatic characterization of some oral and nonoral gram-negative bacteria with the API ZYM system, J. Clin. Microbiol., 14, 288, 1981. 43. Suido, H. et al., Arylaminopeptidase activities of oral bacteria, J. Dent. Res., 65, 1335, 1986. 44. Suido, H. et al., Identification of periodontopathic bacteria based upon their peptidase activities, Adv. Dent. Res., 2, 304, 1988. 45. Gazi, M.I. et al., Comparison of host tissue and bacterial dipeptidylpeptidases in human gingival crevicular fluid by analytical isoelectric focusing, Arch. Oral Biol., 40, 731, 1995. 46. Gazi, M.I. et al., Characterization of protease activities in Capnocytophaga spp., Porphyromonas gingivalis, Prevotella spp., Treponema denticola and Actinobacillus actinomycetemcomitans, Oral Microbiol. Immunol., 12, 240, 1997. 47. Leung, K.P. et al., Surface appendages, hemagglutination, and adherence to human epithelial cells of Bacteroides intermedius, Oral Microbiol. Immunol., 4, 204, 1989.

Molecular Detection of Human Bacterial Pathogens 48. Dorn, B.R, Leung, K.L., and Progulske-Fox, A., Invasion of human oral epithelial cells by Prevotella intermedia, Infect. Immun. 66, 6054, 1998. 49. Taylor-Robinson, D. et al., Oro-dental bacteria in various atherosclerotic arteries, Eur. J. Clin. Microbiol. Infect. Dis., 21,755, 2002. 50. Fiehn, N.E. et al., Identification of periodontal pathogens in atherosclerotic vessels, J. Periodontol., 75, 731, 2005. 51. Gaetti-Jardim Jr., E. et al., Quantitative detection of periodontopathic bacteria in atherosclerotic plaques from coronary arteries, J. Med. Microbiol., 58, 1568, 2009. 52. Okamoto, M. et al., Hemolytic and hemagglutinating activities of Prevotella intermedia and Prevotella nigrescens, FEMS Microbiol. Lett., 178, 299, 1999. 53. Haraldsson, G. et al., Properties of hemagglutination by Prevotella melaninogenica, Anaerobe, 11, 285, 2005. 54. Tamura, M. et al., Effects of zinc and copper on adhesion and hemagglutination of Prevotella intermedia and Prevotella nigrescens, Oral Microbiol. Immunol., 20, 339, 2005. 55. Whittaker, C.J., Klier, C.M., and Kolenbrander, P.E., Mechanisms of adhesion by oral bacteria, Annu. Rev. Microbiol., 50, 513, 1996. 56. Lévesque, C., Lamothe, J., and Frenette, M., Coaggregation of Streptococcus salivarius with periodontopathogens: Evidence for involvement of fimbriae in the interaction with Prevotella intermedia, Oral Microbiol. Immunol., 18, 333, 2003. 57. Kamaguchi, A. et al., Adhesins encoded by the gingipain genes of Porphyromonas gingivalis are responsible for coaggregation with Prevotella intermedia, Microbiology, 149, 1257, 2003. 58. Stefanopoulos, P.K., and Kolokotronis, A.E., The clinical significance of anaerobic bacteria in acute orofacial odontogenic infections, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endodontol., 98, 398, 2004. 59. Siqueira, Jr., J.F., and Rôças, I.N., Bacterial pathogenesis and mediators in apical periodontitis, Braz. Dent. J., 18, 267, 2007. 60. Saito, K. et al., Effects of glucose on formation of cytotoxic end-products and proteolytic activity of Prevotella intermedia, Prevotella nigrescens and Porphyromonas gingivalis, J. Periodontol. Res., 36, 355, 2001. 61. Santos, S.G. et al., The influence of molecular oxygen exposure on the biology of Prevotella intermedia, with emphasis on its antibiotic susceptibility, J. Appl. Microbiol., 103, 882, 2007. 62. Noack, B. et al., Periodontal infections contribute to elevated systemic C-reactive protein level, J. Periodontol., 72, 1221, 2001. 63. Kim, S-J. et al., Prevotella intermedia lipopolysaccharide stimulates release of tumor necrosis factor-α through mitogen-activated protein kinase signaling pathways in monocytederived macrophages, FEMS Immunol. Med. Microbiol., 51, 407, 2007. 64. Chung, Y-H. et al., Lipopolysaccharide from Prevotella nigrescens stimulates osteoclastogenesis in cocultures of boné marrow mononuclear cells and primary osteoblasts, J. Periodontol. Res., 41, 288, 2006. 65. Strömbeck, L. et al., Prevotella bivia can invade human cervix epithelial (HeLa) cells, APMIS, 115, 241, 2007. 66. Pucar, A. et al., Correlation between atherosclerotic and periodontal putative pathogenic bacterial infections in coronary and internal mammary arteries, J. Periodontol., 78, 677, 2007. 67. Love, R.M., Hemin nutritional stress inhibits bacterial invasion of radicular dentine by two endodontic anaerobes, Int. Endodontol. J., 40, 94, 2007.

Prevotella 68. Silva, T.A et al., In vitro activation of the hemolysin in Prevotella nigrescens ATCC 33563 and Prevotella intermedia ATCC 25611, Res. Microbiol., 155, 31, 2004. 69. Guan, S-M. et al., Purification and characterization of a hemoglobin-binding outer membrane protein of Prevotella intermedia, FEMS Microbiol. Lett., 235, 333, 2004. 70. Guan, S.M. et al., Degradation of human hemoglobin by Prevotella intermedia, Anaerobe, 12, 279, 2006. 71. Mallorquí-Fernández, N. et al., A new autocatalytic activation mechanism for cysteine proteases revealed by Prevotella intermedia interpain A, J. Biol. Chem. 283, 2871, 2008. 72. Frandsen, E.V., Kjeldsen, M., and Kilian, M., Inhibition of Prevotella and Capnocytophaga immunoglobulin A1 proteases by human serum, Clin. Diagn. Lab. Immunol., 4, 458, 1997. 73. Yanagisawa, M. et al., Proteinase activity of Prevotella species associated with oral purulent infection, Curr. Microbiol., 52, 375, 2006. 74. Guan, S-M. et al., Prevotella intermedia induces metalloproteinase-9 expression in human periodontal ligament cells, FEMS Microbiol. Lett., 283, 47, 2008. 75. Itoh, T. et al., The activation of matrix metalloproteinases by a whole-cell extract from Prevotella nigrescens, J. Endodontol., 35, 55, 2009. 76. Griswold, K.E., White, B.A., and Mackie, R.I., Diversity of extracellular proteolytic activities among Prevotella species from the rumen, Curr. Microbiol., 39, 187, 1999. 77. Hecht, D.W., Anaerobes: Antibiotic resistant, clinical significance, and the role of susceptibility testing, Anaerobe, 12, 115, 2006. 78. Mosca, A. et al., Antimicrobial profiles of Prevotella spp. and Fusobacterium nucleatum isolated from periodontal infections in a selected area of southern Italy, Int. J. Antimicrob. Agents, 30, 521, 2007. 79. Iwahara, K. et al., Detection of cfxA and cfxA2, the β-lactamase genes of Prevotella spp., in clinical samples from dentoalveolar infection by real-time PCR, J. Clin. Microbiol., 44, 172, 2006. 80. Huits, R.M.H.G. et al., Prevotella bivia necrobacillosis following infectious mononucleosis, J. Infect., 53, e53, 2006.

597 81. Stoakes, L. et al., Gas-liquid chromatographic analysis of cellular fatty acids for identification of Gram-negative anaerobic bacilli, J. Clin. Microbiol., 29, 2636, 1991. 82. Ashimoto, A. et al., Polymerase chain reaction detection of 8 putative periodontal pathogens in subgingival plaque of gingivitis and advanced periodontitis lesions, Oral Microbiol. Immunol., 11, 266, 1996. 83. Martin, F.E. et al., Quantitative microbiological study of human carious dentine by culture and real-time PCR: Association of anaerobes with histopathological changes in chronic pulpitis, J. Clin. Microbiol., 40, 1698, 2002. 84. Kuboniwa, M. et al., Quantitative detection of periodontal pathogens using real-time polymerase chain reaction with TaqMan probes, Oral Microbiol. Immunol., 19, 168, 2004. 85. Yoshida, A. et al., Multiplex polymerase chain reaction assay for simultaneous detection of black-pigmented Prevotella species in oral specimens, Oral Microbiol. Immunol., 20, 43, 2005. 86. Riggio, M.P., and Lennon, A., Development of a novel PCR assay for detection of Prevotella oris in clinical specimens, FEMS Microbiol. Lett., 276, 123, 2007. 87. Frandsen, E.V., Poulsen, K., and Kilian, M., Confirmation of the species Prevotella intermedia and Prevotella nigrescens, Int. J. Syst. Bacteriol., 45, 429, 1995. 88. Teanpaisan, R. et al., Clonality of Porphyromonas gingivalis, Prevotella intermedia and Prevotella nigrescens isolated from periodontally diseased and healthy sites, J. Periodontol. Res., 31, 423, 1996. 89. Milsom, S.E. et al., Rapid differentiation of Prevotella intermedia and P. nigrescens by 16S rDNA PCR-RFLP, J. Med. Microbiol., 44, 41, 1996. 90. Okabe, S. et al., Quantification of host-specific BacteroidesPrevotella 16S rRNA genetic markers for assessment of fecal pollution in freshwater, Appl. Microbiol. Biotechnol., 74, 890, 2007. 91. Nagashima, S. et al., Use of the genomic subtractive hybridization technique to develop a real-time PCR assay for quantitative detection of Prevotella spp. in oral biofilm samples, J. Clin. Microbiol., 43, 2948, 2005. 92. Dahlén, G. et al., A comparison of two transport media for saliva and subgingival samples, Oral Microbiol. Immunol., 8, 375, 1993.

Section IV Proteobacteria Alphaproteobacteria

52 Anaplasma Marina E. Eremeeva and Gregory A. Dasch CONTENTS 52.1 Introduction...................................................................................................................................................................... 601 52.1.1 Classification and Morphology............................................................................................................................. 601 52.1.2 Biology, Habitat, Ecology, and Epidemiology...................................................................................................... 601 52.1.3 Genetic and Genomic Features............................................................................................................................. 603 52.1.4 Clinical Features and Pathogenesis...................................................................................................................... 604 52.1.5 Diagnosis.............................................................................................................................................................. 605 52.1.5.1 Examination of Peripheral Blood Smears............................................................................................. 605 52.1.5.2 Immunohistochemistry.......................................................................................................................... 605 52.1.5.3 Amplification of A. phagocytophilum DNA.......................................................................................... 605 52.1.5.4 In Vitro Cultivation of A. phagocytophilum.......................................................................................... 606 52.1.5.5 Serologic Diagnosis............................................................................................................................... 607 52.2 Methods............................................................................................................................................................................ 607 52.2.1 Sample Preparation............................................................................................................................................... 607 52.2.2 Detection Procedures............................................................................................................................................ 608 52.3 Conclusion and Future Perspectives................................................................................................................................. 608 Acknowledgments.......................................................................................................................................................................612 References...................................................................................................................................................................................612

52.1  INTRODUCTION 52.1.1  Classification and Morphology The genus Anaplasma encompasses several obligately intracellular bacterial species (e.g., Anaplasma phagocytophilum, A. marginale, A. centrale, A. bovis and A. platys) that are classified in the family Anaplasmataceae, order Rickettsiales. While Anaplasma phagocytophilum infects the neutrophils of ruminants (previously referred to as Ehrlichia phagocytophila), equines (Ehrlichia equi), and humans (HGE agent) (Figure 52.1);2 A. marginale, A. centrale, A. bovis and A. platys are essentially animal pathogens, which are beyond the scope of this chapter. The type strain of A. phagocytophilum is strain Webster. Different isolates of A. phagocytophilum are causative agents of tick-borne fever in sheep, cattle, and goats; equine granulocytic anaplasmosis; canine granulocytic anaplasmosis; and human granulocytic anaplasmosis. They exhibit a variety of specific features and host adaptations, which have been summarized in several reviews.2–7 For the purpose of the current review, the scope of this chapter is limited to description of the agent of human granulocytic anaplasmosis (HGA) and the clinical characteristics of this disease. Human granulocytic anaplasmosis was discovered in 1994 in the United States and was originally described as “human granulocytic ehrlichiosis.”8 Since then, numerous reviews

and original articles have been published on this disease, which may be consulted for additional details.9–16 Anaplasma phagocytophilum grows as a cluster of small cocci or pleomorphic coccobacilli in the cytoplasm of neutrophils. Although they are gram-negative, the bacteria do not stain well with the Gram stain and are better visua­lized using Wright or Giemsa staining. Within infected cells, A. phagocytophilum is found inside intracytoplasmic vacuoles, where they form macrocolonies of 1.5–6 µm diameter called morulae.17,18 Two morphological types of individual bacteria are present in morulae, a larger reticulate form and a smaller electron dense form (Figure 52.2); both types of cells divide by binary fission. The bacterium has a typical gram-negative cell wall ultrastructure, containing membrane proteins; however, the cell wall does not contain significant amount of peptidoglycan, and are lacking the lipopolysacharides, pili, and capsule.17,18

52.1.2  Biology, Habitat, Ecology, and Epidemiology Anaplasma phagocytophilum has unusual capacity for infecting and surviving within the vacuole of neutrophils, which they enter via receptor-mediated endocytosis.19–21 Binding of A. phagocytophilum to human neutrophils is mediated at least partially by the adherence of its 44-kDa major surface prtotein-2 (Msp2) to P-selectin glycoprotein ligand,19,22 whereas interaction with murine neutrophils requires 601

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Molecular Detection of Human Bacterial Pathogens

Anaplasma marginale AF414873

100

92 Anaplasma phagocytophilum AY055469 Anaplasma platys M82801 87 Anaplasma bovis U03775 Ehrlichia chaffeensis AF416764

100 84 100

99

Ehrlichia ewingii U96436 Ehrlichia ruminantium CR925677

98

95

Ehrlichia muris EMU15527 Ehrlichia canis CP000107

Candidatus Neoehrlichia mikurensis AB196305 Wolbachia pipientis AF179630 Neorickettsia helminthoeca CP001431 100 100

100

Neorickettsia risticii CP001431 100 Neorickettsia sennetsu CP000237 Rickettsia rickettsii L36217 Rickettsia prowazekii M21789 Orientia tsutsugamushi D38622 Escherichia coli J01859

0.02

FIGURE 52.1  Reconstruction of the phylogenetic position of Anaplasma phagocytophilum. Phylogenetic tree constructed using nucleotide sequences from 16S rRNA gene of Ehrlichia and Anaplasma species and related bacteria, a homologous sequence of E. coli was used as an outgroup. Sequences were ClustalW aligned and tree was drawn using the MEGA4.0 software.1 Distance matrix was calculated using Kimura-2 parameters and tree was obtained using the Neighbor-Joining method based on comparison of 1320 sites after complete deletion of gaps. The scale represents 0.02% divergence. Numbers at nodes are the proportions of 1000 bootstrap resamplings that support the topology shown. Only bootstrap values greater than 80 are shown. The sequences of the isolates used in this analysis are from the NCBI GenBank and the accession numbers are shown next to the sequence name.

FIGURE 52.2  Ultrastructure of A. phagocytophilum in HL-60 cells. Scale bar = 1 μm. The cytoplasm contains several morulae of different size; morulae are shown with black arrows, morula with single reticulate body is shown with an open arrow. (Courtesy of V. Popov. With permission.)

Anaplasma

expression of 1–3-fucosyltransferase.23 Internalized A. phagocytophilum delays apoptosis of infected neutrophils,24 and survives this superoxide-rich environment by scavenging O2−. Once infected by A. phagocytophilum, the modified endosome fails to fuse with lysosomal vesicles and form mature acidified phagolysosomes. A. phagocytophilum exploits host cell nutrients and divides until the cell is lysed, so it can infect other neutrophils. Beside alteration of phagocytic pathways, A. phagocytophilum reduces neutrophil adhesion to endothelial cells; upregulates production of chemokines, mediating transmigration of lymphocytes; and disrupts their motility, degranulation, and respiratory burst.20,25 Anaplasma phagocytophilum is a tick-borne agent that is naturally maintained in a zoonotic cycle between the ticks of the Ixodes persulcatus complex and mammalian reservoir hosts.4,6,13,26 The ecology of A. phagocytophilum is complex. In the United States, it is found in Ixodes scapularis in the eastern and midwestern states, and in I. pacificus and I. spinipalpis in western coastal and mountain areas, respectively.16 The prevalence of A. phagocytophilum in I. scapularis ranges from 4.9% to >50%.27–30 Ixodes persulcatus, I. ovatus, and I.  ricinus are vectors transmitting A. phagocytophilum in Asia and Europe.26,31–36 DNA of A. phagocytophilum has been detected in 2%–45% of I. ricinus ticks (reviewed by Ref. 26); its presence varies from 2.1% to 4.5% in I. persulcatus from different parts of Russia37 to 4%–4.3% in I. persulcatus from China and Korea.32,36,38 In Japan, different genotypes of A. phagocytophilum were detected in 9.6% of I. persulcatus and 6.3% of I. ovatus.35 Many species of domestic and wild animals are important reservoirs of A. phagocytophilum in European and Asian countries. Beside horses, cattle, sheep, and dogs, potential reservoirs include wood mouse (Apodemus sylvaticus), bank vole (Clethrionymus graleolus), shrew (Sorex aranus), roe deer (Capreolus capreolus) and other cervids; foxes (Vulpes vulpes), wild boar (Sus scrofa), bear, and rabbits.33,39–42 The bacterium has also been found in I. trianguliceps from the UK and I. ventalloi from Portugal43–46; the role of these ticks in the natural cycle and transmission of the HGA agent needs further evaluation, but they are not known to feed on humans. In the United States, the white-footed mouse, Peromyscus leucopus, is a primary mammalian reservoir for A. phagocytophilum; however, other small rodents from the genera Neotoma, Apodemus, Microtus, and Clethrionomys are likely reservoirs for A. phagocytophilum and hosts for its tick vector.13,47 Other potential reservoirs in the United States include white-tailed deer (Odocoileus virginianus), raccoons (Procyon lotor), opossums (Didelphis virginiana), squirrels (Sciurus spp.), striped skunks (Mephitis mephitis), chipmunks (Tamias), rabbits, and lizards.48–54 Some of the strains of A. phagocytophilum found in association with different mammalian species are believed to be nonpathogenic for humans, such as Ap-1 variant, which circulates in white-tailed deer.55–57 It is commonly accepted that immature and adult ticks acquire A. phagocytophilum while feeding on bacteremic animals and efficiently pass it transtadially to the next life stage. In contrast, transovarial transmission from an adult

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female tick to its progeny is considered rare.13 However, recent data indicate that transovarial transmission of an A. phagocytophilum variant may occur in Dermacentor albipictus.58 These observations suggest that some types of A. phagocytophilum do not exclusively depend on mammalian reservoirs and horizontal transmission during every generation of ticks. Humans acquire infection as a result of exposures to infected ticks in a variety of habitats. Most infections occur during the spring to summer months, correlating with the biological cycles of feeding of overwintering, infected adult ticks and nymphs infected as larvae. Human anaplasmosis is reportable in the United States59,60; passively collected surveillance data show a steady, increasing trend in the number of annual cases, which reached 1026 in 2008. It is believed that the actual incidence and distribution are significantly greater than is reported.10 Active surveillance estimated from 24 to 58 cases per 100,000 of population (0.02%–0.06%) in Connecticut61 and the Upper Midwest.62 The background seroprevalence rate was approximately 14.9% among healthy individuals from northwestern Wisconsin who had no known history of recent tick bite.63 HGA is not formally reportable outside the United States; therefore, extrapolations regarding the incidence and prevalence depend upon the number of laboratory-confirmed cases reported in the literature. These appear to be infrequent, or the disease is often undetected, since less than 100 laboratoryconfirmed or probable cases of HGA have been reported from Europe and Russia.26,64,65 The temporal distribution and demographic characteristics of the populations affected are similar in the United States and Europe. It has been noticed that a significant discrepancy exists between the high levels of seropositivity to A. phagocytophilum observed in populations from some European countries (up to 17.1%) and the relatively low level of A. phagocytophilum DNA detected in I. ricinus (0.25%–11.5%) in some regions. This suggests that people are frequently exposed to infected ticks and that they may exhibit persistent antibody responses to nonpathogenic strains of A. phagocytophilum found in I.  ricinus.66,67 Seroprevalence to A. phagocytophilum ranges from 8.8% to 20% in different regions of China68,69; antibodies to A.  phagocytophilum were detected by IFA (geometric mean titer 1/747) in 1.8% (n = 491) of febrile patients from Korea.70 HGA can also be acquired rarely through exposure to A. phagocytophilum by blood transfusion, organ transplantation, nosocomial and perinatal transmission, or inhalation of aerosolized blood.71–75

52.1.3  Genetic and Genomic Features The complete genome sequence has been determined for A.  phagocytophilum strain HZ, isolated in 1995 from a patient from New York, USA.76 The genome consists of a single circular chromosome of 1,471,282 bp (NCBI GenBank accession number CP000235) with 41% G + C content. The published annotation predicts 1264 proteins and 42 RNA genes. The gene order of A. phagocytophilum has significant synteny with the genomes of Ehrlichia species. The genome of A.  phagocytophilum contains 462 open reading frames

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(ORFs) or paralog clusters that are unique; the majority of these unique genes encode hypothetical, conserved hypothetical, and conserved domain proteins, as well as uncharacterized membrane proteins and lipoproteins. Other A.  phagocytophilum-specific genes include those enco­ding the P44 outer membrane proteins and the HGE-14 and HGE-2 antigenic proteins. The tick-borne Anaplasmataceae (Ehrlichia spp. and Anaplasma spp.) may be the only Rickettsiales that are not transmitted transovarially in the invertebrate host. One ortholog cluster containing a class II aldolase/adducing domain protein (NSE_0849, RC0678, RP493, RT0479, WD0208) is absent only from Ehrlichia spp. and Anaplasma spp. It has been suggested that a lack of this aldolase/adducing domain protein may prevent transovarial transmission of these agents in the arthropod vector.76 The superoxide dismutase gene, sodB is cotranscribed with components of the type IV secretion system in A. phagocytophilum suggesting that it has an important role in pathogenesis beyond elimination of superoxide. The A. phagocytophilum genome has three omp-1, one msp2, two msp2 homologs, one msp4, and 113 p44 loci belonging to the OMP-1/MSP2/P44 superfamily. Microarray-based comparative genomic hybridization reveals that expansion of the p44 family is a common feature in A. phagocytophilum strains.76 The genome of A.  phagocytophilum has numerous genes encoding HGE-14 protein that may be an excreted effector molecule of its type IV secretion system. A. phagocytophilum has genetic capacity to synthesize all nucleotides, most vitamins, and cofactors. A complete pyruvate dehydrogenase, tricarboxylic acid cycle, F0F1-ATPase, and electron transport chain were found. It appears to use host-derived carboxylates and amino acids but does not obtain carbon or energy from fatty acids or actively carry out glycolysis. Differential gene transcription profiles were demonstrated for isolate HZ grown in HL-60, human microvascular endothelial cells (HMEC-1), and a tick cell line.77 In particular, it was shown that seven virB2 paralogs of the type IV secretion system exhibited human or tick cell dependent transcription. Unique cell-type dependent expression of previously unrecognized genes and coding sequences were identified, including expression of p44/msp2 paralogs (of 114 total, only two are detected in HL-60, three in HMEC-1, and none in tick cells) suggesting variable synthesis of these proteins during different stages of the A. phagocytophilum life cycle.

52.1.4  Clinical Features and Pathogenesis HGA is an acute illness that occurs 5–21 days after the bite of an infected tick14; its severity ranges from asymptomatic seroconversion to a mild or severe febrile illness. Most patients experience fever >101°F, myalgia, headache, and malaise.14,62 Other symptoms occur less frequently and include nausea, abdominal pain, vomiting, diarrhea, cough, and arthralgias. Involvement of the skin or central nervous system is very rare during the course of HGA. Basic laboratory findings included thrombocytopenia (45 days) is definitively required.

54.1.4  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 spillover of brucellosis from wildlife to livestock and the globalization of animal trade promote the worldwide distribution of animal and human disease.2,61 Hence, despite being controlled in many developed countries, brucellosis remains a major public health concern. Bovine brucellosis (B. abortus) 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.”62 Norway and Switzerland are also considered to be brucellosis-free. In contrast, the situation is less favorable in southern European countries.63 Ovine/caprine brucellosis (B. melitensis) 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 (B. suis) occurs in most areas in which pigs are kept. The human pathogenic biovars B. suis bv 1 and bv 3 are mainly isolated from wild boars, feral swine, and domestic pigs in Asia, South America, in the southeastern states of the United States, and in Queensland, Australia.1,61 Available epidemiological evidence shows that B. suis bv 2 is the most common agent of porcine brucellosis in Europe,

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but the biovars 1 and 3 also occur.64 B. suis bv 2 has only exceptionally been described as the causative agent of human brucellosis.65,66 The epidemiology of human brucellosis has significantly changed in the last decade for sanitary, socioeconomic, and political reasons. Currently, about a half million new cases are annually reported worldwide, and prevalence rates exceed 10 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.67,68 Although ovine/caprine brucellosis has a limited geographic distribution, B. melitensis is by far the main cause of clinically apparent disease in humans worldwide.69,70 In territories where ovine/caprine brucellosis is endemic and the population density of sheep and goats is high, there is a significant relationship between the number of infected flocks and the incidence of human cases.71 B. abortus and B. suis infections are often associated with certain occupational groups, for example, farm workers, veterinarians, and meatpacking employees, whereas human B.  melitensis infections frequently occur without occupational exposure.46 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 the B. abortus infections most frequently reported before 1960 to the B. suis infections predominating in the early 1970s and, currently, to an increased reporting of B. melitensis infections.72 In nonendemic countries, an association of human brucellosis with the immigrant population has been described.73–75 Most cases have been imported, and the disease was acquired either through international travel or contaminated food products from endemic areas.76 In the United States, 50%–60% of the brucellosis cases are notified in California and Texas, mainly among Hispanics.67,77 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.67,72,75,77

54.1.5  M  icrobiological and Serological Methods in Diagnosis of Brucellosis 54.1.5.1  Isolation and Identification Comparable to other bacterial infections, blood culture is still the gold standard in the diagnosis of human brucellosis. Although blood is the material most commonly examined, bone marrow cultures may yield a higher sensitivity, especially in patients with previous antibiotic treatment, and culture times may be faster.78 Samples of any body fluid and of tissue collected during surgery or at necropsy can also be cultured. The isolation rate of brucellae from specimens with a competing microflora or from pus is low. In patients suffering from acute brucellosis the sensitivity of blood cultures

Molecular Detection of Human Bacterial Pathogens

may vary from 80% to 90% whereas in chronic disease direct bacteriologic confirmation is less successful ranging between 30% and 70% depending on the method of isolation used.79 Brucella grows on most standard media, for example, blood agar, chocolate agar, trypticase soy agar (TSA), or serum-dextrose agar (SDA). Two to five percent 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°C–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.80 As the number of Brucella organisms is likely to be low in tissue samples, enrichment culture using liquid media may be helpful. Castañeda’s medium, a nonselective, biphasic medium, has been recommended for the isolation of Brucella from clinical samples.81,82 However, weekly subcultures onto solid selective medium for up to 6 weeks are necessary. Therefore, this traditional method was replaced by automated culture systems, such as the lysis centrifugation method, which helped to reduce culture times and increased sensitivity.83,84 After a 2- to 3-day incubation period, punctate, 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-Sserum) mixed with a saline suspension of colonies. 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 spp. and biovars 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, for example, as Moraxella phenylpyruvica.85 Since drug resistance does not significantly contribute to treatment failures and relapses, in vitro susceptibility testing of Brucella isolates is not generally recommended.86 Brucella strains recovered from relapsed patients usually show antibiotic sensitivity profiles identical to the strains recovered during primary infection.86,87

Brucella

54.1.5.2  Serological Tests Because of the shortcomings of culture techniques, which are time consuming, hazardous, and not sensitive, most physicians rely on the indirect proof of Brucella infections. The detection of high or rising titers of specific antibodies in the serum allows a tentative diagnosis. IgM antibodies are predominant in the first week after inoculation with Brucella.88 A switch to IgG isotype antibodies can be observed in the second week, followed by a continuous rise in both IgM and IgG levels, reaching a peak about four weeks after infection. Antibody titers usually decline with the beginning of an adequate antibiotic treatment, but significant antibody titers may persist for up to a year despite successful therapy.88,89 IgG antibodies are related to active infection, and they are constantly detected in bacteriologically proven cases. Chronic courses of the disease are highly unlikely when IgG antibodies cannot be detected anymore.88 However, antibody titers can peak a second time (usually only IgG), which might be a sign of relapse or an incomplete recovery.90 In summary, the evaluation of Brucella-specific IgM and IgG antibodies helps to discriminate patients suffering from brucellosis at an early stage of disease or acute brucellosis from those with chronic brucellosis.91 The serological diagnosis of brucellosis is traditionally based on agglutination tests, for example, the serum agglutination test (SAT); the Rose Bengal test (RBT); and the antiglobulin, or Coombs’, test (CT), which are all used for screening. The SAT developed by Wright and Smith in 1897 is still the most popular serological method in the diagnosis of human brucellosis.92 In addition, SAT is frequently used as a reference to which other serological tests are compared.93 Particularly the quality of the antigen preparations is essential for consistent and reproducible results in agglutination tests. Nevertheless, cross-reactions with various bacteria, for example, Yersinia enterocolitica O:9, Escherichia coli O:157, Francisella tularensis, Salmonella urbana group N, Vibrio cholerae, and Stenotrophomonas maltophilia, have been reported.93 A confirmatory test is therefore always recommended to avoid false-positive results due to the low specificity of agglutination tests. Samples that reacted positive in a screening test are retested using the complement fixation test (CFT) or an enzyme-linked immunosorbent assay (ELISA), both of which are suitable for screening and confirmation. The sensitivity of various serological tests might also be low because they are based on secondary interactions such as the ability to agglutinate antigens or to fix complement. Especially in the early stage of infection, false-negative or only weak positive reactions may occur. Furthermore, the interpretation of test results is subjective reflecting the experience of the investigator. Both, sensitivity and specificity of agglutination tests for brucellosis depend on the cut-off value used and on the background level of reactive antibodies in the population.79 Therefore, cut-off values for agglutination titers considered to be confirmatory for active disease cannot be easily defined. In general, a titer ≥1:160 provides strong supportive

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evidence for active infection, and titers ≥1:80 are supposed to be suspicious, but even lower titers are not unusual in active brucellosis.79,93 Especially in endemic areas, intermediate titers are difficult to interpret. In urban populations, a titer of 1:80 can be considered positive; whereas, in rural areas, where brucellosis is endemic, titers of ≥ 1:320 might be linked to disease.94 Early in the course of disease or in persistent brucellosis, incomplete or blocking (nonagglutinating) antibodies may lead to negative serological test results even in patients with positive blood culture.95 Last but not least, the sensitivity of serological tests can be increased by testing paired serum samples. When testing a follow-up sample collected a few weeks after the initial diagnosis, seroconversion or a significant increase in antibody titers may prove active infection. Conventional Serological Tests for Antibody Detection. Serum Agglutination Test: Several methods are suitable for agglutination tests, for example, the tube agglutination test (Wright test), slide or plate agglutination tests, and the card agglutination test. In the classical tube agglutination test, serial dilutions of the serum to be tested are mixed with the antigen, for example, a standardized phenol-inactivated suspension of a smooth culture of B. abortus organisms (strain 1119-3) adjusted to pH 7.2–7.4. The mixture is incubated in a water bath at 37°C for 48 h, and finally, the test results are recorded as grade of agglutination. The serum titer is defined as the dilution in which at least 50% of the bacterial cells are clumped. The tube agglutination test is labor intensive and time consuming. Rose Bengal Test: The RBT was developed in the United States as a rapid screening test in the 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 ± 0.05.96 Acidification loosens the bond between the antigen and nonspecific agglutinins, leading to an improved specificity and sensitivity of the RBT in comparison with SAT.97,98 Buffered plate antigen tests have proven to be inexpensive screening tests in humans with reduced nonspecific reactions in comparison with the SAT.99 However, in endemic areas the use of the RBT as the sole technique for the diagnosis of brucellosis has to be considered very carefully, particularly in patients repeatedly exposed to Brucella or with a history of brucellosis.100 Complement Fixation Test: 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. Especially during the incubation period and in the late chronic stage of disease, when SAT results are either negative or inconclusive, CFT has proven to be the diagnostic tool of choice. Primary Binding Assays for Antibody Detection. Enzyme-Linked Immunosorbent Assay (ELISA): A great variety of antigens has been used for indirect enzyme-linked immunosorbent assay (iELISA), including whole cells, sLPS, native hapten polysaccharide, salt-extracted proteins, outer membrane proteins, and cytoplasmic proteins.93 However,

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most ELISAs are based on bacterial extracts including high amounts of LPS, and false-positive reactions can occur after exposure to cross-reacting bacteria. Competitive enzyme immunoassays (cELISAs) are less prone to cross-reacting antibodies than conventional tests or iELISAs. Because of their additional high sensitivity, cELISAs can be used as confirmatory and screening tests. ELISA is an excellent method for screening large populations for the presence of anti-Brucella antibodies.101 In addition, ELISA techniques are helpful in classifying the stage of disease according to the detected antibody profile.102 Patients suffering from acute brucellosis usually show very high levels of specific IgG and significant levels of IgM and IgA, whereas patients with chronic brucellosis show moderately high levels of IgG. Low levels of IgG in healthy persons may represent residual antibodies even a long time after complete recovery from brucellosis. Since the performance of ELISAs depends on the stage of disease, IgM- and IgG-ELISAs should generally be performed in parallel to increase sensitivity.103–105 Fluorescence Polarization Assay (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.106 FPA can be performed within a few minutes and requires minimal sample preparation. Under field conditions even whole blood samples can be analyzed.107 For culture­confirmed human brucellosis cases the sensitivity of the FPA is 96%, and the specificity determined in healthy blood donors is 98%.106 Lateral Flow Assay (LFA): LFA can be used as a bedside test for the serodiagnosis of human brucellosis at all stages of disease.108,109 For test performance, only a drop of blood obtained by a finger prick is needed. Furthermore, the LFA does not require a specific hands-on training, and test results are easy to interpret. All components, that is, the B. abortus antigen spotted onto a nitrocellulose strip and the monoclonal antihuman antibodies conjugated to colloidal dye particles for nonenzymatic detection, are stabilized and do not require cooling either during transport or for storage. Sensitivity and specificity of the LFA are high (>95%). Hence, LFA is a suitable rapid point-of-care assay ideal for field testing in an outbreak setting and for screening large populations.

54.1.6  Molecular Diagnostic Techniques 54.1.6.1  PCR Numerous PCR techniques have been developed for the identification of Brucella.110 PCR methods are increasingly used in the diagnosis of human 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. PCR turned out to be a suitable

Molecular Detection of Human Bacterial Pathogens

tool for the detection of Brucella in various matrices, including serum, whole blood and urine samples, different tissues, cerebrospinal fluid, synovial or pleural fluid, and pus.111–115 Currently, the preferred clinical specimens used for molecular diagnosis of human brucellosis are whole blood and serum samples. Whether the serum fraction is superior to whole blood for molecular detection of the pathogen, or vice versa, is controversially discussed.116,117 Although the concentration of PCR inhibitors is lower in serum samples, the small number of bacteria circulating after treatment may result in the absence of the target DNA in serum leading to false-negative results. For rapid diagnosis of human brucellosis, the simple identification of Brucella by genus-specific PCR is adequate because antibiotic treatment is independent of the disease‑causing species. A lot of target sequences have been used for detection of the genus Brucella, for example, the 16S rRNA,118,119 the 16S-23S internal transcribed spacer region (ITS),120 a gene coding for an outer membrane protein (omp2),121 and the insertion element IS711.122,123 However, the majority of genus-specific PCR assays target the bcsp31 gene, which codes for a 31-kDa immunogenic outer membrane protein conserved among all Brucella spp.124 For species-specific surveillance programs, differential PCR assays are needed. The multiplex AMOS–PCR using a five primer cocktail was a first approach toward the differentiation of four Brucella species, that is, B. abortus (bvs 1, 2, and 4); B. melitensis (bvs 1, 2, and 3); B. ovis; and B. suis (bv 1).125 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 of this insertion element, leading to variable amplicon sizes. Recently, a conventional multiplex PCR assay suitable for the identification of all Brucella species (except for B. inopinata) and the vaccine strains, B. abortus RB51, B. abortus S19, and B. melitensis Rev1 has been developed.126 The identification is based on the numbers and sizes of seven products amplified by the so-called Bruce-ladder PCR (Figure 54.1). In a worldwide multicenter study, the Bruce-ladder PCR has proven to be a useful tool for the rapid identification of Brucella strains in basic microbiology laboratories.127 Another conventional multiplex PCR, based on primers targeting specific deletions and insertions in the different Brucella genomes, partially allows subtyping of Brucella species at the biovar level.128 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.129,130 Test results are available within a few hours. The species-specific real-time PCRs established by Redkar et al.,131 Newby et al.,132 and Probert et al.133 were described as rapid, sensitive, and specific diagnostic tools with a low risk of cross-contamination and the potential of automation. Single nucleotide polymorphism (SNP)-based differentiation of Brucella clades incorporated into real-time PCR assays may also provide a rapid method for the identification of Brucella species.134

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B. microti

B. ceti

B. pinnipedialis

B. neotomae

B. canis

B. ovis

B. melitensis

B. suis

B. abortus

Brucella

FIGURE 54.1  Identification and differentiation of Brucella species by the Bruce-ladder PCR.126,127

Various studies of genus-specific real-time PCR assays applied to human specimens have been published. A realtime PCR targeting the bcsp31 gene appears to be suitable for screening whenever Brucella is suspected because false-negative results caused by rare species and biovars may be avoided by a genus-specific approach. Consecutively, a species-specific assay should be applied because the use of more than one marker-based PCR may increase sensitivity and specificity.130 Since PCR reactions may be inhibited by clinical samples,135 an internal amplification control should be included.130 Dilution of the samples may also reduce inhibitory effects of the sample matrix but the generally small amount of target 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 sensitivity. Since the number of bacteria in the clinical samples of patients suffering from focal forms of brucellosis is usually small, the detection capacity of a valuable molecular assay has to be very high. Sensitivity of conventional and real-time PCR assays for the diagnosis of brucellosis can be further increased using the IS711 insertion element of Brucella as a target sequence because it is found in multiple copies in Brucella chromosomes.136,137 Definite criteria to establish the success of treatment are still missing, and to date, none of the conventional microbiological methods has proven to be efficient to predict relapse. In the last years, more studies have been conducted to evaluate the usefulness of PCR methods during the incubation period and for posttreatment follow-up of Brucella infections. Quantitative real-time PCR appears to be a useful method to identify symptomatic nonfocal disease in patients for whom the classical microbiological methods fail.138 In addition, real-time PCR using serum samples is a valuable tool, not

only for the initial diagnosis of brucellosis, but also for the differentiation between active and past infection.139 Using single-step conventional PCRs for posttreatment follow-up in whole blood samples, PCR results were negative in successfully treated patients, whereas test results were positive in relapsed patients.116,140 In contrast, using real-time PCR techniques, Brucella DNA could be detected in the majority of brucellosis patients throughout treatment and follow-up, despite appropriate antibiotic therapy and apparent clinical recovery.141,142 Although DNA load constantly decreases after the end of treatment, a significant number of patients exhibit bacterial DNA load even years after clinical cure and in the absence of any symptoms indicative of disease persistence or relapse.142 However, Brucella may persist in human macrophages and replicate in such a low frequency that transient bacteremia can be controlled by the immune system. Furthermore, the diagnostic yield of modern realtime PCR assays is better as compared to conventional techniques leading to the detection of nonviable or phagocytosed microorganisms. No difference in the evolution of DNA load can be observed between patients who relapse and those who do not.143 Nevertheless, quantitative real-time PCR could be helpful in the diagnosis of chronic infection. B. melitensis DNA can be detected in asymptomatic subjects with a history of brucellosis, albeit in a smaller proportion than in the group of symptomatic patients suffering from chronic disease.138 The clinical significance of persisting bacterial DNA load in asymptomatic patients long after successful completion of antibiotic therapy is still unclear. Further studies are urgently needed to clarify whether the persistence of Brucella DNA in human serum reflects the presence of active bacteria possibly leading to recurrent disease or just nonviable bacterial fragments. Real-time PCR methods can also be useful in the differential diagnosis of brucellosis. For instance, in areas of high incidence of brucellosis and tuberculosis, differential diagnosis is often difficult because both diseases are characterized by clinical polymorphism and granoulomatous inflammation. In addition, Brucella spp. and Mycobacterium tuberculosis are slow-growing pathogens, and culture may be insensitive in extrapulmonary tuberculosis and focal manifestations of brucellosis. For rapid differential diagnosis a multiplex realtime PCR assay based on the intergenic region SenX3-ReX3 and bcsp31 has recently been developed.115 54.1.6.2  Molecular Typing Methods Molecular epidemiology significantly contributes to the analysis and understanding of infections caused by pathogenic bacteria.144 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 it spreads. Fast and accurate typing procedures are therefore crucial for the eradication and control of brucellosis. However, molecular typing is difficult due to the high degree of genetic homology (>90%) among Brucella species, demonstrated by DNA–DNA hybridization,25 due to identical 16S rRNA gene sequences among all Brucella species,145 and because more than 90% of

636

all genes share >98% sequence identity.23 Nevertheless, various methods have been established for molecular subtyping of Brucella strains, for example, AP–PCR (arbitrary primed– PCR),146 ERIC–PCR (enterobacterial repetitive intergenic consensus sequence–PCR), REP–PCR (repetitive intergenic palindromic sequence–PCR),147,148 RAPD–PCR (random amplified polymorphic DNA–PCR),149 PCR–RFLP (PCR– restriction fragment length polymorphism) of different genetic loci,150 AFLP (amplified fragment length polymorphism),151 SNP (single nucleotide polymorphism),152,153 and MLST (multilocus sequence typing).154 However, with the exception of SNP and MLST, these tests lack reproducibility, show a limited capability to differentiate single strains, and are not appropriate for routine typing. Based on the genomes of B. melitensis 16M,21 B. suis 1330,22 and B. abortus strain 9-94123 tandem repeat loci with a high degree of size polymorphism and variable number tandem repeats (VNTRs) could be identified. 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. Tandem repeats are classified in satellites (megabases of DNA) present in many eukaryotic genomes, minisatellites (spanning hundreds of base pairs, with a repeat unit size of at least 9 bp), and microsatellites (spanning a few tens of nucleotides with a repeat unit size up to 8 bp).155 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 discriminatory among unrelated Brucella isolates that could not be differentiated by classical microbiological methods. The hypervariability found by MLVA is in contrast to the wellknown genetic homogeneity of the genus. However, tandem repeats generally mutate at different rates.155 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.156 The hypermutability of VNTR loci has been exploited for Brucella in large panels of animal isolates.157–160 Various VNTR-based genotyping methods have been tested for subtyping human Brucella isolates both with a broad global presentation156 and from small spatial scales, such as the Middle East,161 Lebanon,162 Italy,163 Portugal,164 Mexico,165 and Peru.166,167 Currently, the most frequently used molecular typing assay is the MLVA-16 scheme, which includes eight minisatellite loci (panel 1) and eight microsatellites (panel 2, which is subdivided into panels 2A and 2B) (Table 54.4).156 The MLVA-16 data can be queried on the genotyping Web page at http://mlva.u-psud.fr/. MLVA-16 has also proven to be useful for clinical applications. MLVA-16 allows identifying the source of laboratoryacquired Brucella infections168 and helps to differentiate relapse from reinfection, which may have implications on therapy.87 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

Molecular Detection of Human Bacterial Pathogens

(Figure 54.2) or automatic high-throughput procedures, that is, an automated capillary electrophoresis-based method161 or a lab on a chip technology.169 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 that fulfils all performance criteria recommended for a typing assay, that is, typeability, reproducibility, stability, discriminatory power, concordance with other typing techniques, and epidemiological concordance.159

54.2  METHODS 54.2.1  Sample Collection and Preparation Blood, urine, cerebrospinal fluid, synovial fluid, and tissue can be valuable specimens for direct detection of Brucella DNA.115 Due to the small number of brucellae in these clinical specimens, special methods for DNA preparation are required to reduce inhibitory effects caused by matrix components and to concentrate the DNA. The preparation of a bacterial DNA template from serum simply by boiling does not prevent the presence of PCR inhibitors.135 Commercial kits such as the QIAampTM DNA Mini Kit (Qiagen Inc., Valencia, California) have been successfully used to extract Brucella DNA from whole blood, serum, and tissue samples.142,143,170 Queipo-Ortuño 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, California).171 However, comprehensive evaluation studies on DNA extraction methods in different human samples are still missing.

54.2.2  Detection Procedures 54.2.2.1  Multiplex PCR and MLVA The previously described AMOS–PCR is highly effective for the differentiation of B. abortus, B. melitensis, B. ovis, and B. suis125 (Table 54.1). This PCR employs one primer

TABLE 54.1 Primers for AMOS–PCR Primers BA BM BS BO IS711

Sequence (5′→3′) GAC GAA CGG AAT TTT TCC AAT CCC AAA TCG CGT CCT TGC TGG TCT GA GCG CGG TTT TCT GAA GGT TCA GG CGG GTT CTG GCA CCA TCG TCG TGC CGA TCA CTT AAG GGC CTT CAT

Source: Bricker, B.J., and Halling, S.M., J. Clin. Microbiol., 11, 2660, 1994.

637

Brucella

(IS711) for binding with the insertion sequence IS711, which is present in all Brucella species, and four primers for recognizing the species-specific regions downstream of this insertion sequence, leading to formation of variable sizes of amplicons from these four Brucella species, and facilitating their identification. To identify all Brucella species (except for B. inopinata) and the vaccine strains B. abortus RB51, B. abortus S19, and B. melitensis Rev1, the Bruce-ladder PCR can be applied.126,127 Use of primers and PCR conditions shown in Tables 54.2 and 54.3, respectively, will result in the amplification of distinct bands from most Brucella species (Figure 54.1).

TABLE 54.2 Primers for Bruce-Ladder PCR Primers

Amplicon Size (bp)

Sequence (5′→3′)

BMEI0998f BMEI0997r BMEI0535f

ATC CTA TTG CCC CGA TAA GG GCT TCG CAT TTT CAC TGT AGC GCG CAT TCT TCG GTT ATG AA

BMEI0536r

CGC AGG CGA AAA CAG CTA TAA

BMEII0843f BMEII0844r BMEI1436f BMEI1435r BMEII0428f BMEII0428r BR0953f BR0953r BMEI0752f BMEI0752r BMEII0987f BMEII0987r Bmispec f Bmispec r

TTT ACA CAG GCA ATC CAG CA GCG TCC AGT TGT TGT TGA TG ACG CAG ACG ACC TTC GGT AT TTT ATC CAT CGC CCT GTC AC GCC GCT ATT ATG TGG ACT GG AAT GAC TTC ACG GTC GTT CG GGA ACA CTA CGC CAC CTT GT GAT GGA GCA AAC GCT GAA G CAG GCA AAC CCT CAG AAG C GAT GTG GTA ACG CAC ACC AA CGC AGA CAG TGA CCA TCA AA GTA TTC AGC CCC CGT TAC CT AGA TAC TGG AAC ATA GCC CG ATA CTC AGG CAG GAT ACC GC

1682 450 (1320 in Brucella strains isolated from marine mammals) 1071 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. Source: García-Yoldi, D. et al., Clin. Chem., 52, 779, 2006; López-Gõni, I. et al., J. Clin. Microbiol. 46, 3484, 2008.

TABLE 54.3 Bruce-Ladder PCR Cycling Program Initial denaturation Denaturation Annealing Extension Last extension

Temperature (°C)

Time

95 94 58 72 72

15 min 30 s 90 s 120 s 10 min

Cycles 1 25

1

The recently developed MLVA-16 assay156 has proved to be highly discriminatory among unrelated human Brucella isolates that could not be differentiated by other PCR methods or by the classical microbiological techniques. The MLVA-16 is based on two panels of primers (Table 54.4), one comprising eight minisatellite markers (panel 1) useful for species identification, and a second group of eight complementary microsatellite markers (panel 2) showing a higher discriminatory power useful for molecular epidemiology. The panel 2 markers are split into two groups, panel 2A and 2B, composed of three and five markers, respectively. Panel 2B contains the most highly variable markers. 54.2.2.2  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 54.5 and 54.6).130,133 Using IS711 as target the sensitivity of real-time PCR technology can be further increased because this intergenic element occurs in multiple copies within the Brucella genome (Table 54.7). Assays for the species-specific identification of B. melitensis described by Redkar et al. and Probert et al. have a low detection limit (Table 54.8).130,131,133 Assays designed for the detection of B. abortus by Redkar et al., Probert et al., and Newby et al. only detected biovars 1, 2, and 4, because of the more distant relatedness of other biovars (Table 54.9).131–133 A  real-time PCR assay specific for B. suis bv 1 was also established by Redkar et al.131 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 can be easily transferred from one technical platform to another,172 diagnostic laboratories are able to choose between hybridization probes technology and 5′ exonuclease assays to meet 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 Tables 54.5 through 54.9.

54.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES In the diagnosis of human brucellosis, time-consuming culture and phenotypic characterization of the isolate is still the “gold standard.” The low yield of Brucella cultures, however, often results in diagnostic delay and the late initiation of appropriate antibiotic therapy. Serology is a more effective means of diagnostic, although cross-reactivity is still a major problem. PCR has proven to be a valuable tool when culture fails or serological results are inconclusive, and 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

638

Molecular Detection of Human Bacterial Pathogens

TABLE 54.4 Primers for MLVA-16 Primer (5′→3′) VNTR Locus

Upper

Lower

Panel 1 bruce06-RU1322_134bp_408bp_3u bruce08-BRU1134_18bp_348bp_4u bruce11-BRU211_63bp_257bp_2u bruce12-BRU73_15bp_392bp_13u bruce42-BRU424_125bp_539bp_4u bruce43-BRU379_12bp_182bp_2u bruce45-BRU233_18bp_151bp_3u bruce55-BRU2066_40bp_273bp_3u

ATGGGATGTGGTAGGGTAATCG ATTATTCGCAGGCTCGTGATTC CTGTTGATCTGACCTTGCAACC CGGTAAATCAATTGTCCCATGA CATCGCCTCAACTATACCGTCA TCTCAAGCCCGATATGGAGAAT ATCCTTGCCTCTCCCTACCAG TCAGGCTGTTTCGTCATGTCTT

GCGTGACAATCGACTTTTTGTC ACAGAAGGTTTTCCAGCTCGTC CCAGACAACAACCTACGTCCTG GCCCAAGTTCAACAGGAGTTTC ACCGCAAAATTTACGCATCG TATTTTCCGCCTGCCCATAAAC CGGGTAAATATCAATGGCTTGG AATCTGGCGTTCGAGTTGTTCT

Panel 2A bruce18-BRU339_8bp_146bp_5u bruce19-Bru324_6bp_163bp_18u bruce21-BRU329_8bp_148bp_6u

TATGTTAGGGCAATAGGGCAGT GACGACCCGGACCATGTCT CTCATGCGCAACCAAAACA

GATGGTTGAGAGCATTGTGAAG ACTTCACCGTAACGTCGTGGAT GATCTCGTGGTCGATAATCTCATT

Panel 2B bruce04-BRU1543_8bp_152bp_2u bruce07-BRU1250_8bp_158bp_5u bruce09-BRU588_8bp_156bp_7u bruce16-BRU548_8bp_152bp_3u bruce30-BRU1505_8bp_151bp_6u

CTGACGAAGGGAAGGCAATAAG GCTGACGGGGAAGAACATCTAT GCGGATTCGTTCTTCAGTTATC ACGGGAGTTTTTGTTGCTCAAT TGACCGCAAAACCATATCCTTC

CGATCTGGAGATTATCGGGAAG ACCCTTTTTCAGTCAAGGCAAA GGGAGTATGTTTTGGTTGTACATAG GGCCATGTTTCCGTTGATTTAT TATGTGCAGAGCTTCATGTTCG

Source: Al Dahouk, S. et al., J. Microbiol. Methods, 69, 137, 2007.

a

b

c

d

e

f

FIGURE 54.2  MLVA typing of Brucella field isolates of various geographic origins using panel 1 VNTR markers.156 (a and b) Turkey, (c) Italy, (d) Pakistan, (e) United Arab Emirates, (f) Syria.

and probes should always be included in a diagnostic scheme to detect infrequently isolated Brucella species and atypical strains. 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 patients is low.

There has been controversy about the preferred clinical specimen, the best method of DNA extraction, and the PCR assays with the lowest limit of detection. Most real-time PCR assays developed for the detection of Brucella can detect fewer than 10 bacteria per reaction, which is close to the technical limit of this method. In addition, a variety of DNA purification methods have been validated that eliminate

639

Brucella

TABLE 54.5 Real-Time PCR Assay Targeting bcsp31 for Specific Detection of Genus Brucella A. PCR Mixture Makeup

Brucella spp. (bcsp31)

Reagent Water 25 mM MgCl2 10× concentrated Reaction Mix B4 B5 Bru FL Bru LC Lambda F Lambda R Lambda FL Lambda LC Phage Lambda DNAa Sample DNA a

Stock Concentration (pmol/µL = µM)

LightCycler FastStart DNA Master Hybridization Probes Cat. No. 2 239 272 Primers and Probes (5′→3′) 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

2.5 mM

20 pmol/µL 20 pmol/µL 8 µM 8 µM 20 pmol/µL

0.5 0.5 0.5 0.5 0.5

0.5 µM 0.5 µM 0.2 µM 0.2 µM 0.5 µM

20 pmol/µL 8 µM 8 µM

0.5 0.5 0.5 1 2

0.5 µM 0.15 µM 0.15 µM

Inclusion of an internal amplification control is especially useful for clinical samples.130

B. PCR Parameter Parameter Activation of Fast Start Taq DNA polymerase Amplification (45 cycles)

Melting curve analysis

Cooling

Temperature (°C)

Time

95

10:00 min

Slope (°C/s) Acquisition Mode 20

None

95 55 72 95 55 95 40

10 s 10 s 12 s 0 30 s 0 30 s

20 20 20 20 20 0.1 20

None Single None None None Continuous None

Note: The assay specifically detects all brucellae but no other organisms, with a detection limit of 16 fg DNA in buffer.

inhibitory components of the matrix. Remaining limitations are the small number of brucellae to be found in clinical specimens and, consequently, the even smaller amount of target DNA available for PCR. Despite of these biological and technical problems, especially for real-time PCR methods, various advantages compared to standard serological and bacteriological methods could have been demonstrated. However, PCR results are not always reproducible in different laboratories. PCR assays targeting different genes may help to exclude false-positive results. In order to determine inhibitory effects of the matrix, an internal amplification control has to be integrated into all PCR 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 the context of other diagnostic tests and of course the clinical signs and symptoms of the patient. Molecular techniques may revolutionize the diagnosis of human brucellosis. Real-time PCR technology meets all requirements for a rapid diagnosis in clinical microbiology laboratories. The drastic reduction of diagnostic delay will have important prognostic implications in life-threatening complications of the disease, such as Brucella endocarditis or neurobrucellosis.

640

Molecular Detection of Human Bacterial Pathogens

TABLE 54.6 Real-Time 5′-Exonuclease Assay Targeting bcsp31 for Large-Scale Screening of Genus Brucella A. PCR Mixture Makeup Stock Concentration (pmol/µL = µM)

Brucella spp. (bcsp31)

Reagent Water Reaction buffer

Final Concentration (in 25µL)

µL Per Reaction   5.5

2×

12.5

1×

Brucella spp fw

TaqMan Universal MasterMix (uMM) Applied Biosystems Cat. No. 4304447 Primers and Probes (5′→3′) 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

50 95 95 57

2 min 10 min 15 s 1 min

 1  1 45

Decontamination Initial denaturation Denaturation Annealing

Note: 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 54.7 Real-Time PCR Assay Targeting Intergenic Element IS711 for Specific and More Sensitive Detection of Genus Brucella A. PCR Mixture Makeup

Reagent Water 25 mM MgCl2 10× concentrated Reaction Mix

Brucella spp. (IS711)

Stock Concentration (pmol/µL = µM)

µL Per Reaction 12.4     1.6

Final Concentration (in 20 µL) 3 mM

IS711_S

LightCycler FastStart DNA Master Hybridization Probes Cat. No. 2 239 272 Primers and Probes (5′→3′) TTGTCGATGCTATCGGCCTAC

20 pmol/µL

0.5

0.5 µM

IS711_R

GGCAATGAAGGCCCTTAAGT

20 pmol/µL

0.5

0.5 µM

IS711_FL

GAAGCTTGCGGACAGTCACCATAAT--FL

8 µM

0.5

0.2 µM

IS711_LC

LC Red640-GCCGGGTGTTGGCTTTATTCG--PH

8 µM

0.5

0.2 µM

Sample DNA

2

2 continued

641

Brucella

TABLE 54.7 (continued) Real-Time PCR Assay Targeting Intergenic Element IS711 for Specific and More Sensitive Detection of Genus Brucella B. PCR Parameter Parameter

Temperature (°C)

Time

Slope (°C/s)

95

10:00 min

20

None

95 55 72 95 45 95 40

10 s 10 s 15 s 0 30 s 0 30 s

20 20 20 20 20   0.1 20

None Single None None None Continuous None

Activation of Fast Start Taq DNA polymerase Amplification (45 cycles)

Melting curve analysis

Cooling

Acquisition Mode

TABLE 54.8 Real-Time PCR Assays Targeting BMEI1162/IS711 for Specific Detection of B. melitensis Designed as 5′-Exonuclease Assay for Large-Scale Screening A. PCR Mixture Makeup

Brucella melitensis BMEI1162/IS711

Reagent Water Reaction buffer

B. melitensis fw B. melitensis rev B. melitensis Taq

Stock Concentration (pmol/µL = µM)

TaqMan Universal MasterMix (uMM) Applied Biosystems Primers and Probes (5′→3′) AACAAGCGGCACCCCTAAAA CATGCGCTATGATCTGGTTACG 6FAM-CAGGAGTGTTTCGGCTCAGAATAATC CACA-DB

Sample DNA

µL Per Reaction

Final Concentration (in 25µL)

2×

  5.5 12.5

1×

10 µM 10 µM 10 µM

  0.75   0.75   0.5

0.3 µM 0.3 µM 0.2 µM

 5

B. PCR Parameter Decontamination Initial denaturation Denaturation Annealing

Temperature (°C) 50 95 95 57

Time 2 min 10 min 15 s 1 min

Cycle  1  1 45

Note: This assay specifically detects all B. melitensis isolates but no other orgnisms with a detection limit of 16fg DNA in buffer. Source: Probert, W.S. et al., J. Clin. Microbiol., 42, 1290, 2004.

642

Molecular Detection of Human Bacterial Pathogens

TABLE 54.9 Real-Time PCR Assay Targeting alkB/IS711 for Specific Detection of B. abortus Designed as 5′-Exonuclease Assay for Large-Scale Screening A. PCR Mixture Makeup

Reagent

Stock Concentration (pmol/µL = µM)

Brucella abortus alkB/IS711

Water

µL Per Reaction

Final Concentration (in 25 µL)

  5.5

Reaction buffer

TaqMan Universal MasterMix (uMM) Applied 2× Biosystems Primers and Probes (5′→3′)

12.5

1x

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-CGCTCATGCTCGCCAG ACTTCAATG-DB

10 µM

  0.5

0.2 µM

Sample DNA

 5

B. PCR Parameter Decontamination Initial denaturation Denaturation Annealing

Temperature (°C)

Time

Cycle

50 95 95 57

2 min 10 min 15 s 1 min

 1  1 45

Note: 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.

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 Young, E.J., and Corbel, M.J. (Editors), Brucellosis: Clinical and Laboratory Aspects, p. 53, CRC-Press, Florida, 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. 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 Krieg, N.R., and Holt, J.G. (Editors), Bergey’s Manual of Systematic Bacteriology, vol. 1, p. 370, Williams & Wilkins, Baltimore, 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. Hubalek, Z. et al., Brucellosis of the common vole (Microtus arvalis), Vector Borne Zoonotic Dis., 7, 679, 2007. 14. Scholz, H.C. et al., Brucella microti sp. nov., isolated from the common vole Microtus arvalis, Int. J. Syst. Evol. Microbiol., 58, 375, 2008. 15. Scholz, H.C. et al., Isolation of Brucella microti from mandibular lymph nodes of red foxes, Vulpes vulpes, in Lower Austria, Vector Borne Zoonotic Dis., 9, 153, 2009. 16. Scholz, H.C. et al., Isolation of Brucella microti from soil, Emerg. Infect. Dis., 14, 1316, 2008. 17. De, B.K. et al., Novel Brucella strain (BO1) associated with a prosthetic breast implant infection, J. Clin. Microbiol., 46, 43, 2008. 18. Scholz, H.C. et al., Brucella inopinata sp. nov, isolated from a breast implant infection, Int. J. Syst. Evol. Microbiol., 60, 801, 2010.

Brucella 19. Gee, J.E. et al., Use of 16S rRNA gene sequencing for rapid confirmatory identification of Brucella isolates, J. Clin. Microbiol., 42, 3649, 2004. 20. Scholz, H.C. et al., Genetic diversity and phylogenetic relationships of bacteria belonging to the Ochrobactrum-Brucella group by recA and 16S rRNA gene-based comparative sequence analysis, Syst. Appl. Microbiol., 31, 1, 2008. 21. DelVecchio, V.G. et al., The genome sequence of the facultative intracellular pathogen Brucella melitensis, Proc. Natl. Acad. Sci. USA, 99, 443, 2002. 22. 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. 23. 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. 24. 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. 25. Verger, J.M. et al., Brucella, a monospecific genus as shown by deoxyribonucleic acid hybridization, Int. J. Syst. Bacteriol., 35, 292, 1985. 26. 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. 27. Moreno, E., and Moriyón, I., Brucella melitensis: a nasty bug with hidden credentials for virulence, Proc. Natl. Acad. Sci. USA, 99, 1, 2002. 28. Lapaque, N. et al., Brucella lipopolysaccharide acts as a virulence factor, Curr. Opin. Microbiol., 8, 60, 2005. 29. 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. 30. 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. 31. Roop II, R.M. et al., Adaptation of the brucellae to their intracellular niche, Mol. Microbiol., 52, 621, 2004. 32. Boschiroli, M.L. et al., The Brucella suis VirB operon is induced intracellularly in macrophages, Proc. Natl. Acad. Sci. USA, 99, 1544, 2002. 33. Celli, J., and Gorvel, J.P., Organelle robbery: Brucella interactions with the endoplasmic reticulum, Curr. Opin. Microbiol., 7, 1, 2004. 34. 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. 35. Köhler, S. et al., What is the nature of the replicative niche of a stealthy bug named Brucella? Trends Microbiol., 11, 215, 2003. 36. Al Dahouk, S. et al., Proteomic analysis of Brucella suis under oxygen deficiency reveals flexibility in adaptive expression of various pathways, Proteomics, 9, 3011, 2009. 37. 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. 38. Miller, M.A., and Paige, J.C., Other food borne infections, Vet. Clin. North Am. Food Anim. Pract., 14, 71, 1998.

643 39. 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. 40. Brew, S.D. et al., Human exposure to Brucella recovered from a sea mammal, Vet. Rec., 144, 483, 1999. 41. 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. 42. Sohn, A.H. et al., Human neurobrucellosis with intracerebral granuloma caused by a marine mammal Brucella sp., Emerg. Infect. Dis., 9, 485, 2003. 43. 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. 44. Yagupsky, P., and Baron, E.J., Laboratory exposures to brucellae and implications for bioterrorism, Emerg. Infect. Dis., 8, 1180, 2005. 45. 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. 46. Young, E.J., An overview of human brucellosis, Clin. Infect. Dis., 21, 283, 1995. 47. Colmenero, J.D. et al., Complications associated with Brucella melitensis infection: A study of 530 cases, Medicine (Baltimore), 75, 195, 1996. 48. 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. 49. 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. 50. Food and Agriculture Organization-World Health Organiza­ tion, Joint FAO/WHO Expert Committee on Brucellosis (fifth report), in WHO Technical Report Series, No. 464, p.  34, WHO, Geneva, 1971. 51. Food and Agriculture Organization-World Health Organiza­ tion, Joint FAO/WHO Expert Committee on Brucellosis (sixth report), in WHO Technical Report Series, No. 740, p. 56, World Health Organization, Geneva, 1986. 52. Hall, W.H., Modern chemotherapy for brucellosis in humans, Rev. Infect. Dis., 12, 1060, 1990. 53. Solera, J., Martinez-Alfaro, E., and Espinosa, A., Recognition and optimum treatment of brucellosis, Drugs, 53, 246, 1997. 54. Solera, J., Treatment of human brucellosis, J. Med. Liban., 48, 255, 2000. 55. Corbel, M.J. et al., Treatment of human brucellosis. In Corbel, M.J., Elberg, S.S., and Cosivi, O. (Editors), Brucellosis in Humans and Animals, Chapter 5, World Health Organization, Geneva, 2006. 56. Ariza, J. et al., Comparative trial of rifampin-doxycycline versus tetracycline-streptomycin in the therapy of human brucellosis, Antimicrob. Agents Chemother., 28, 548, 1985. 57. Ariza, J. et al., Treatment of human brucellosis with doxycycline plus rifampin or doxycycline plus streptomycin. A randomized, double-blind study, Ann. Intern. Med., 117, 25, 1992. 58. Solera, J. et al., Doxycycline-rifampin versus doxycyclinestreptomycin in treatment of human brucellosis due to Brucella melitensis. The GECMEI Group. Grupo de Estudio de Castilla-la Mancha de Enfermedades Infecciosas, Antimicrob. Agents Chemother., 39, 2061, 1995.

644 59. Colmenero, J.D. et al., Comparative trial of doxycycline plus streptomycin versus doxycycline plus rifampin for the therapy of human brucellosis, Chemotherapy, 35, 146, 1989. 60. Solera, J. et al., Treatment of human brucellosis with doxycycline and gentamicin, Antimicrob. Agents Chemother., 41, 80, 1997. 61. Godfroid, J., Brucellosis in wildlife, Rev. Sci. Tech., 21, 277, 2002. 62. 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. 63. Taleski, V. et al., An overview of the epidemiology and epizootology of brucellosis in selected countries of Central and Southeast Europe, Vet. Microbiol., 90, 147, 2002. 64. European Food Safety Authority (EFSA), Scientific opinion of the panel on animal health and welfare (AHAW) on a request of the commission on porcine brucellosis (Brucella suis), EFSA J., 1144, 1, 2009. 65. Paton, N.I. et al., Visceral abscesses due to Brucella suis infection in a retired pig farmer, Clin. Infect. Dis., 32, 129, 2001. 66. Teyssou, R. et al., About a case of human brucellosis due to Brucella suis biovar 2, Mèd. Mal. Infect., 19, 160, 1989. 67. 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. 68. Wise, R.I., Brucellosis in the United States: past, present, and future, JAMA, 244, 2318, 1980. 69. Corbel, M.J., Brucellosis: an overview, Emerg. Infect. Dis., 3, 213, 1997. 70. Pappas, G. et al., Medical Progress—Brucellosis, N. Engl. J. Med., 352, 2325, 2005. 71. De Massis, F. et al., Correlation between animal and human brucellosis in Italy during the period 1997–2002, Clin. Microbiol. Infect., 11, 632, 2005. 72. Fosgate, G.T. et al., Time-space clustering of human brucellosis, California, 1973–1992, Emerg. Infect. Dis., 8, 672, 2002. 73. Al Dahouk, S. et al., Changing epidemiology of human brucellosis, Germany, 1962–2005, Emerg. Infect. Dis., 13, 1895, 2007. 74. Eriksen, N. et al., Brucellosis in immigrants in Denmark, Scand. J. Infect. Dis., 34, 540, 2002. 75. White, Jr., A.C., and Atmar, R.L., Infections in Hispanic immigrants, Clin. Infect. Dis., 34, 1627, 2002. 76. Memish, Z.A., and Balkhy, H.H., Brucellosis and international travel, J. Travel Med., 11, 49, 2004. 77. 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. 78. Gotuzzo, E. et al., An evaluation of diagnostic methods for brucellosis—the value of bone marrow culture, J. Infect. Dis., 153, 122, 1986. 79. Franco, M.P. et al., Human brucellosis, Lancet Infect. Dis., 7, 775, 2007. 80. 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. 81. Castañeda, M.R., A practical method for routine blood cultures in brucellosis, Proc. Soc. Exp. Biol. Med., 64, 114, 1947. 82. Yagupsky, P., Detection of brucellae in blood cultures, J. Clin. Microbiol., 37, 3437, 1999. 83. Bannatyne, R.M., Jackson, M.C., and Memish, Z., Rapid diagnosis of Brucella bacteremia by using the BACTEC 9240 system, J. Clin. Microbiol., 35, 2673, 1997.

Molecular Detection of Human Bacterial Pathogens 84. Mantur, B.G., and Mangalgi, S.S., Evaluation of conventional Castaneda and lysis centrifugation blood culture techniques for diagnosis of human brucellosis, J. Clin. Microbiol., 42, 4327, 2004. 85. Barham, W.B. et al., Misidentification of Brucella species with use of rapid bacterial identification systems, Clin. Infect. Dis., 17, 1068, 1993. 86. Ariza, J. et al., Relevance of in vitro antimicrobial susceptibility of Brucella melitensis to relapse rate in human brucellosis, Antimicrob. Agents Chemother. 30, 958, 1986. 87. Al Dahouk, S. et al., Failure of a short-term antibiotic therapy for human brucellosis using ciprofloxacin—a study on in vitro susceptibility of Brucella strains, Chemotherapy, 51, 352, 2005. 88. Reddin, J.L. et al., Significance of 7S and macroglobulin Brucella agglutinins in human brucellosis, N. Engl. J. Med., 24, 1263, 1965. 89. Buchanan, T.M. et al., Brucellosis in the United States, 1960– 1972: An abattoir-associated disease. II. Diagnostic aspects, Medicine, 53, 415, 1974. 90. Gazapo, E. et al., Changes in IgM and IgG antibody concentrations in brucellosis over time: Importance for diagnosis and follow-up, J. Infect. Dis., 159, 219, 1989. 91. Araj, G.F., Human brucellosis: A classical infectious disease with persistent diagnostic challenges, Clin. Lab. Sci. 12, 207, 1999. 92. Wright, A.E., and Smith, F., On the application of the serum test to the differential diagnosis of typhoid fever and Malta fever, Lancet, 149, 656, 1897. 93. 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. 94. Jumba, M.M., Mirza, N.B., and Mwaura, F.B., Agglutinins for brucellae antigens in blood sera of an urban and rural population in Kenya, East Afr. Med. J., 73, 204, 1996. 95. Young, E.J., Human brucellosis, Rev. Infect. Dis. 5, 821, 1983. 96. Morgan, W.J.B. et al., The rose bengal plate agglutination test in the diagnosis of brucellosis, Vet. Rec., 85, 636, 1969. 97. 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. 98. 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. 99. Lucero, N.E., and Bolpe, J.E., Buffered plate antigen test as a screening test for diagnosis of human brucellosis, J. Clin. Microbiol., 36, 1425, 1998. 100. Ruiz-Mesa, J.D. et al., Rose Bengal test: Diagnostic yield and use for the rapid diagnosis of human brucellosis in emergency departments in endemic areas, Clin. Microbiol. Infect., 11, 221, 2005. 101. Caravano, R., Chabaud, F., and Oberti, J., Application of immunoenzymatic techniques for epidemiological surveys on brucellosis among human populations, Ann. Inst. Pasteur Microbiol., 138, 79, 1987. 102. Magee, J.T., An enzyme-labeled immunosorbent assay for Brucella abortus antibodies, J. Med. Microbiol., 13, 167, 1980. 103. Osoba, A.O. et al., Diagnostic value of Brucella ELISA IgG and IgM in bacteremic and nonbacteremic patients with brucellosis, J. Chemother., 13, 54, 2001. 104. Cakan, G. et al., Assessment of diagnostic enzyme-linked immunosorbent assay kit and serological markers in human brucellosis, Jpn. J. Infect. Dis., 61, 366, 2008.

Brucella 105. Gómez, M.C., et al., Evaluation of seven tests for diagnosis of human brucellosis in an area where the disease is endemic, Clin. Vaccine Immunol., 15, 1031, 2008. 106. Lucero, N.E. et al., Fluorescence polarization assay for diagnosis of human brucellosis, J. Med. Microbiol., 52, 883, 2003. 107. Nielsen, K. et al., Fluorescence polarization assay for the diagnosis of bovine brucellosis: Adaptation to field use, Vet. Microbiol., 80, 163, 2001. 108. Smits, H.L. et al., Development and evaluation of a rapid dipstick assay for serodiagnosis of acute human brucellosis, J. Clin. Microbiol., 37, 4179, 1999. 109. Smits, H.L. et al., Immunochromatographic Brucella-specific immunoglobulin M and G lateral flow assays for rapid serodiagnosis of human brucellosis, Clin. Diagn. Lab. Immunol., 10, 1141, 2003. 110. Bricker, B.J., PCR as a diagnostic tool for brucellosis, Vet. Microbiol., 90, 435, 2002. 111. Colmenero, J.D. et al., Real-time polymerase chain reaction: a new powerful tool for the diagnosis of neurobrucellosis, J. Neurol. Neurosurg. Psychiatry, 76, 1025, 2005. 112. 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. 113. Queipo-Ortuño, 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. 114. Queipo-Ortuño, M.I. et al., Rapid diagnosis of Brucella epididimo-orchitis by real-time polymerase chain reaction assay in urine samples, J. Urol., 176, 2290, 2006. 115. Queipo-Ortuño, M.I. et al., Rapid differential diagnosis between extrapulmonary tuberculosis and focal complications of brucellosis using a multiplex real-time PCR assay, PLoS One, 4, e4526, 2009. 116. Mitka, S. et al., Evaluation of different PCR assays for early detection of acute and relapsing brucellosis in humans in comparison with conventional methods, J. Clin. Microbiol., 45, 1211, 2007. 117. Zerva, L. et al., Serum is the preferred clinical specimen for diagnosis of human brucellosis by PCR, J. Clin Microbiol., 39, 1661, 2001. 118. Herman, L., and De Ridder, H., Identification of Brucella spp. by using the polymerase chain reaction, Appl. Environ. Microbiol., 6, 2099, 1992. 119. Romero, C. et al., Specific detection of Brucella DNA by PCR, J. Clin. Microbiol., 3, 615, 1995. 120. 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., 5, 1683, 1996. 121. 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. 122. 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. 123. 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. 124. Baily, G.G. et al., Detection of Brucella melitensis and Brucella abortus by DNA amplification, J. Trop. Med. Hyg., 95, 271, 1992.

645 125. 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. 126. 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. 127. López-Gõni, I. et al., Evaluation of a multiplex PCR assay (Bruce-ladder) for molecular typing of all Brucella species, including the vaccine strains, J. Clin. Microbiol. 46, 3484, 2008. 128. Huber, B. et al., Development of a PCR assay for typing and subtyping of Brucella species, Int. J. Med. Microbiol., 299, 563, 2009. 129. Al Dahouk, S. et al., The detection of Brucella spp. using PCR-ELISA and real-time PCR assays, Clin. Lab., 50, 387, 2004. 130. 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. 131. Redkar, R. et al., Real-time detection of Brucella abortus, Brucella melitensis and Brucella suis, Mol. Cell. Probes, 15, 43, 2001. 132. 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. 133. 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. 134. Foster, J.T. et al., Real-time PCR assays of single-nucleotide polymorphisms defining the major Brucella clades, J. Clin. Microbiol., 46, 296, 2008. 135. Queipo-Ortuño, M.I. et al., Preparation of bacterial DNA template by boiling and effect of immunoglobulin G as an inhibitor in real-time PCR for serum samples from patients with brucellosis, Clin. Vaccine Immunol., 15, 293, 2008. 136. Bounaadja, L. et al., Real-time PCR for identification of Brucella spp.: A comparative study of IS711, bcsp31 and per target genes, Vet. Microbiol., 137, 156, 2009. 137. Hinic, V. et al., Novel identification and differentiation of Brucella melitensis, B. abortus, B. suis, B. canis, and B. neotomae suitable for both conventional and real-time PCRsystems, J. Microbiol. Methods, 75, 375, 2008. 138. Castaño, M.J., and Solera, J., Chronic brucellosis and persistence of Brucella melitensis DNA, J. Clin. Microbiol., 47, 2084, 2009. 139. Queipo-Ortuño, M.I. et al., Usefulness of a quantitative realtime PCR assay using serum samples to discriminate between inactive, serologically positive and active human brucellosis, Clin. Microbiol. Infect., 14, 1128, 2008. 140. Morata, P. et al., Posttreatment follow-up of brucellosis by PCR assay, J. Clin. Microbiol. 37, 4163, 1999. 141. 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. 142. Vrioni, G. et al., An eternal microbe: Brucella DNA load persists for years after clinical cure, Clin. Infect. Dis., 46, 131, 2008. 143. Maas, K.S. et al., Evaluation of brucellosis by PCR and persistence after treatment in patients returning to the hospital for follow-up, Am. J. Trop. Med. Hyg., 76, 698, 2007.

646 144. van Belkum, A., High-throughput epidemiologic typing in clinical microbiology, Clin. Microbiol. Infect., 9, 86, 2003. 145. Gee, J.E. et al., Use of 16S rRNA gene sequencing for rapid confirmatory identification of Brucella isolates, J. Clin. Microbiol., 42, 3649, 2004. 146. 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. 147. Mercier, E. et al., Polymorphism in Brucella strains detected by studying distribution of two short repetitive DNA elements, J. Clin. Microbiol., 5, 1299, 1996. 148. Tscherneva, E. et al., Repetitive element sequence based polymerase chain reaction for typing of Brucella strains, Vet. Microbiol., 51, 169, 1996. 149. Tscherneva, E. et al., Differentiation of Brucella species by random amplified polymorphic DNA analysis, J. Appl. Microbiol., 88, 69, 2000. 150. 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. 151. 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. 152. Marianelli, C. et al., Molecular characterization of the rpoB gene in Brucella species: New potential molecular markers for genotyping, Microbes Infect., 8, 860, 2006. 153. 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. 154. 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. 155. Vergnaud, G., and Denoeud, F., Minisatellites: Mutability and genome architecture, Genome Res., 10, 899, 2000. 156. Al Dahouk, S. et al., Evaluation of Brucella MLVA typing for human brucellosis, J. Microbiol. Methods, 69, 137, 2007. 157. Le Flèche, P. et al., Evaluation and selection of tandem repeat loci for a Brucella MLVA typing assay, BMC Microbiol., 6, 9, 2006. 158. 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. 159. 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.

Molecular Detection of Human Bacterial Pathogens 160. 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. 161. Tiller, R.V. et al., Comparison of two multiple-locus variablenumber tandem-repeat analysis methods for molecular strain typing of human Brucella melitensis isolates from The Middle East, J. Clin. Microbiol., 47, 2226, 2009. 162. Kattar, M.M. et al., Evaluation of a multilocus variablenumber tandem-repeat analysis scheme for typing human Brucella isolates in a region of brucellosis endemicity, J. Clin. Microbiol., 46, 3935, 2008. 163. Marianelli, C. et al., Molecular epidemiological characterization and antibiotic susceptibility of Brucella isolates from humans in Sicily, Italy, J. Clin. Microbiol., 45, 2923, 2007. 164. Almendra, C. et al., “HOOF-Print” genotyping and haplotype inference discriminates among Brucella spp. isolates from a small spatial scale, Infect. Genet. Evol., 9, 104, 2009. 165. Rees, R.K. et al., Single tube identification and strain typing of Brucella melitensis by multiplex PCR, J. Microbiol. Methods, 78, 66, 2009. 166. Smits, H.L. et al., MLVA genotyping of human Brucella isolates from Peru, Trans. R. Soc. Trop. Med. Hyg., 103, 399, 2009. 167. Nöckler, K. et al., Molecular epidemiology of Brucella genotypes in patients at a major hospital in central Peru, J. Clin. Microbiol., 47, 3147, 2009. 168. Marianelli, C. et al., Use of MLVA-16 to trace the source of a laboratory-acquired Brucella infection, J. Hosp. Infect., 68, 274, 2008. 169. De Santis, R. et al., Lab on a chip genotyping for Brucella spp. based on 15-loci multi locus VNTR analysis, BMC Microbiol., 9, 66, 2009. 170. 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. 171. 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. 172. 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.

55 Ehrlichia Jere W. McBride, Juan P. Olano, and Nahed Ismail CONTENTS 55.1 Introduction...................................................................................................................................................................... 647 55.1.1 Microbiology, Genetics, and Pathobiology........................................................................................................... 647 55.1.1.1 Taxonomy and Etiologic Agents............................................................................................................ 647 55.1.1.2 Morphology............................................................................................................................................ 648 55.1.1.3 Genetic and Antigenic Features............................................................................................................. 648 55.1.1.4 Pathobiology.......................................................................................................................................... 649 55.1.2 Epidemiology, Clinical Features, and Pathogenesis............................................................................................. 650 55.1.2.1 Epidemiology and Public Health........................................................................................................... 650 55.1.2.2 Human Monocytotropic Ehrlichiosis..................................................................................................... 650 55.1.2.3 Human Ehrlichiosis Ewingii.................................................................................................................. 650 55.1.2.4 Clinical Features of Human Ehrlichosis................................................................................................ 650 55.1.2.5 Pathogenesis of Human Ehrlichiosis..................................................................................................... 651 55.1.3 Diagnosis of Human Ehrlichiosis......................................................................................................................... 652 55.1.3.1 Conventional Techniques....................................................................................................................... 652 55.1.3.2 Molecular Techniques............................................................................................................................ 652 55.2 Methods............................................................................................................................................................................ 653 55.2.1 Sample Preparation............................................................................................................................................... 653 55.2.2 Detection Procedures............................................................................................................................................ 653 55.3 Conclusion and Future Perspectives................................................................................................................................. 653 References.................................................................................................................................................................................. 654

55.1  INTRODUCTION 55.1.1  Microbiology, Genetics, and Pathobiology 55.1.1.1  Taxonomy and Etiologic Agents Many organisms in the order Rickettsiales, represented in the genera Ehrlichia, Anaplasma, Cowdria, and Neorickettsia, have recently been realigned based on genetic sequence information from two universally conserved gene sequences and the application of molecular phyologenetics. Previous criteria relied more heavily on morphologic attributes, cellular tropism, and antigenic similarity to define taxonomic position and classify various “rickettsia-like” organisms in the corresponding genera without the benefit of supporting genotypical information. A new taxonomy for the genus Ehrlichia was proposed in 2001 using data obtained from two highly conserved genes, rrs (16S ribosomal RNA genes) and groESL,1 and completed genome sequences from Ehrlichia and Anaplasma spp. supported the taxonomic positions determined by individual gene sequences. The genus Ehrlichia is now part of a newly created family Anaplasmataceae, which also includes the genera Anaplasma, Wolbachia, and Neorickettsia, but it remains in the order Rickettsiales. The

genus has five approved members (E. canis, E. chaffeensis, E. muris, E. ruminantium, and E. ewingii) that have at least 97.7% similarity in the 16S rRNA sequences, with the acquisition of one member from the genus Cowdria (C. ruminantium). Six previously recognized members have been reassigned to the genera, Anaplasma (E. phagocytophila, E. equi, E. platys, and E. bovis) and Neorickettsia (E. sennetsu and E. risticii). There are three recognized human pathogens in the genus, including E. chaffeensis, E. ewingii, and E. canis (Table 55.1), as well as a pathogen strictly of veterinary importance, E. ruminantium. This chapter focuses on the ehrlichial agents associated with human disease; E. chaffeensis; the etiologic agent of human monocytotropic ehrichiosis (HME); E. ewingii, the agent of human ehrlichiosis ewingii (HEE); and E. canis, the primary etiologic agent of canine monocytic ehrichiosis (CME), and most recently associated with human infections in South America.2 E. chaffeensis. E. chaffeensis was isolated in 1991 and formally named after it caused the first human infection in 1986.3,4 E. chaffeensis exhibits tropism for monocytes/macrophages and causes human monocytotropic ehrlichiosis (HME), 647

648

Molecular Detection of Human Bacterial Pathogens

TABLE 55.1 Ehrlichia spp. That Cause Human Disease Agent

Target Cell

E. chaffeensis

Monocyte

E. ewingii

Neutrophil

E. canis

Monocyte

Natural Hosts

Disease

White-tailed deer, Human canines and monocytotropic goats ehrlichiosis (HME) Canines and deer Human ehrlichiosis ewingii (HEE) Canines Canine monocytic ehrlichiosis (CME)

a life-threatening zoonosis that is also associated with mild to severe infections in dogs.5,6 In humans, many (40%–60%) E. chaffeensis infections require hospitalization,7,8 and there is a case fatality rate of 3% due to the difficulty in making an early and accurate diagnosis. E. chaffeensis is maintained in nature in a zoonotic cycle primarily in white-tailed deer, but dogs may also be a significant natural reservoir.9 The primary vector is the lone star tick, Amblyomma americanum, which is distributed from west central Texas throughout the southeastern, south central and mid-Atlantic states.9,10 E. chaffeensis DNA has been detected in other tick species, including Dermacentor variabilis, Ripicephalus sanguineus, Ixodes pacificus, I. rincinus, A. testudinarium, and Haemaphysalis yeni. Larval ticks become infected with E. chaffeensis after feeding on an infected vertebrate hosts and maintain a transstadial but not transovarial infection. The emergence of E. chaffeensis appears to coincide with changes in demographic and ecologic factors including increases in vector and mammalian host populations and human contact with natural foci, immunocompromised and aging human populations, and improved diagnosis and reporting.9 E. chaffeensis can be cultivated in vitro in various mammalian and tick cell lines11 but causes only transient subclinical infection in immunocompetent mice.12 E. ewingii. E. ewingii was described in 1971 as a pathogen of dogs13 and has been associated commonly with two distinct clinical syndromes in canines, anemia, and polyarthritis.14,15 E. ewingii is transmitted by the lone star tick, A. americanum, and exhibits host cell tropism for polymorphonuclear neutrophils.16 The most common clinical features of E. ewingii infection in dogs are fever, lameness, and neurologic abnormalities.17 E. ewingii as a human pathogen emerged in 1996, and four cases of human ehrlichosis ewingii (HEE) in immunocompromised patients were characterized between 1996 and 1998 that were clinically indistinguishable from disease caused by E. chaffeensis.18 The natural host of E. ewingii is canines, and it appears to be an opportunistic pathogen in humans. Most cases of HEE are manifested in immunocompromised patients, such as those infected with the human immunodeficiency virus (HIV).18,19 Many of the documented human cases reported contact with dogs before the onset of symptoms.18 E. chaffeensis and E. canis appear to be more conserved antigenically and have substantial serologic cross reactivity, and E. ewingii appears to be more

antigenically divergent. There are serologic cross-reactive antigens between E. chaffeensis and E. ewingii, associated with higher (>40 kDa) molecular weight proteins, but epitopes in the major outer membrane proteins (OMP-1s/ p28s) appear to be antigenically distinct.18,20 E. ewingii has not been cultivated in vitro; thus molecular characterization of this agent has been limited to a few genes that include the OMP-1/P28 gene families.20 55.1.1.2  Morphology Ehrlichia species are gram-negative bacteria that stain dark blue with Romanovsky‑type stains.21 Ehrlichiae reside in an intracellular vacuole bounded by a host cell–derived membrane-forming inclusions (morulae) that contain variable numbers of bacteria.9 By electron microscopy, individual bacteria within the vacuole appear coccoid to pleomorphic and vary in size from small (0.4 μm), medium (0.7 μm), large (1 μm), and occasionally very large (≤2 μm).22,23 Two distinct morphological forms can be distinguished, dense-cored and reticulate cells, both of which have a gram-negative envelope, including a cytoplasmic membrane and an outer membrane (cell wall), separated by a periplasmic space. In contrast to free-living gram-negative bacteria, peptidoglycan and lipopolysaccharide have not been detected, which can be explained by the recent revelation that genes for the synthesis of these components are not present in the genome.24 The dense-cored cells are usually smaller, and more consistently coccoid in appearance, with an electron-dense nucleoid that occupies most of the cytoplasm. By analogy to chlamydiae, these cells are also often referred to, particularly in the early literature, as elementary bodies. On the other hand, reticulate cells are larger, more pleomorphic, and have a dispersed filamentous nucleoid and ribosomes dispersed in the cytoplasm. Sometimes organisms of intermediate size and electron density are referred to as intermediate bodies.22,23,25,26 Reticulate cells multiply by binary fission and intermediate bodies can also be observed dividing. Evidence suggests that densecored cells are the infectious form of the organism.27,28 The morphology of E. ruminantium in experimentally infected ticks appears to be similar to that in mammalian cells, consisting of both dense-cored and reticulate forms.29 55.1.1.3  Genetic and Antigenic Features The obligate intracellular lifestyle of Ehrlichia spp. has resulted in genomes that have evolved differently than freeliving microbes. The genomes of E. chaffeensis, E. canis, and E. ruminantium are completed,30–32 but the genome of E. ewingii has not yet been sequenced. Some of the common overall features of the completed genomes include a high degree of gene order, and all are relatively small (1.2, 1.3, and 1.5 Mbp, respectively) due to massive loss of biosynthetic genes through a process known as reductive evolution. Each genome has roughly 1000 protein-encoding genes and a consistently low average G + C content of only 27%–29%. These genomes have a substantially lower than average ratio of coding to noncoding sequence (62%–72% coding) and

Ehrlichia

contain pseudogenes (17–32).30,32 The genomes of these ehrlichiae exhibit a serine-threonine bias in proteins associated with host-pathogen interactions, proteins with tandem and/or ankyrin repeats, protein secretion systems, a large group of secreted proteins with transmembrane helicies suggestive of a complex membrane structure and paralogous protein families that may be involved in immune evasion.30,32 The evolution of Ehrlichia spp. as obligate intracellular organisms has resulted in loss of many metabolic pathway genes. Genes essential for the glycolytic pathway, ATP/ADP translocases are absent, thus they are unable to utilize glucose as a carbon or energy source. In contrast to Rickettsia spp. that are not confined to a vacuole, Ehrlichia spp. have complete pathways for the synthesis of purines and pyrimidines. Genes for enzymes involved in aerobic respiration, sets of genes involved in synthesis of lipids and phospholipids, and have tRNAs for all amino acids.30,32 In addition, they lack genes for the biosynthesis of lipopolysaccharide and peptidoglycan.24 Molecular characterizations of some Ehrlichia spp. genes were independently identified and utilized for initial molecular characterizations and comparisons, including universally conserved genes, rrs, rrl, rrf (16S, 23S, and 5S rRNA, respectively),33–35 groEL, and groESL,36–38 quinolate synthetase A (nadA)39 and citrate synthase (gltA).40 Others involved in pathobiology have been examined for functional roles including thio-disulfide oxidoreductase (dsb),41 ferric-ion binding protein (fbp),42 a virB operon (type IV secretion machinery),43 and two-component regulatory systems.44,45 The interaction of Ehrlichia with the host immune response has been examined using Western immunoblotting, and a small group of E. chaffeensis, E. canis, and E. ewingii proteins that are strongly recognized by antibody have been identified. Major immunoreactive E. chaffeensis proteins observed by immunoblotting include, 200-, 120-, 88-, 55-, 47-, 40-, 28-, and 23-kDa46,47; E. canis, 200-, 140-, 95-, 75-, 47-, 36-, 28-, and 19-kDa48; and E. ruminantium, 160-, 85-, 58-, 46-, 40-, 32-, and 21-kDa.49 E. ewingii has not been cultured, and immunoreactive proteins are not well defined; however, sera from E. ewingii infected dogs consistently cross-react with high (>40 kDa) molecular mass E. canis and E. chaffeensis proteins but not with low molecular mass (2:1) patients >40 years of age; the majority (>80%) report a tick bite,8,95 and HME outbreaks are associated with recreational or occupational activities.90,112 Tick attachment has to last for 24–48 h before disease can occur. HME presents as a more severe disease in patients older than 60 years of age and in immunocompromised patients, including HIV/AIDS patients in whom severe complications, such as adult respiratory distress syndrome, acute tubular necrosis, shock, and CNS involvement, can arise. Another risk factor for severe disease in

651

Ehrlichia

HIV/AIDS patients is sulfa intake, which has been reported to aggravate ehrlichiosis. HME and HEE manifest as undifferentiated febrile illnesses 1–3 weeks after the bite of an infected tick. For HME, the most frequent clinical findings reported anytime during acute illness are fever, malaise, headache, dizziness, chills, and myalgias7,8,95,113–116 (see Table 55.2 for a comprehensive description of clinical manifestations of HME). Patients with HEE present with a milder disease with few complications.18 A vast majority of these patients are immunocompromised, further suggesting that E. ewingii is less pathogenic. Hematologic and biochemical abnormalities usually include leukopenia, thrombocytopenia, anemia, mildly elevated serum hepatic transaminase activities, and hyponatremia.8,19,95 Lymphocytosis characterized by a predominance of γδ T cells is often seen in patients during recovery.117 A high proportion of immunocompetent (41%–62%) and immunocompromised patients (86%) require hospitalization,8,19,95 and delays in antibiotic treatment are associated with more pulmonary complications, increased transfer to intensive care, and longer length of illness.118 The case fatality rate for HME is estimated to be 3%,93 and fatal disease is most often described

in older patients and patients debilitated by underlying disease or immunodeficiency.7,19 Immunocompromised patients (HIV-infected persons, transplant recipients, corticosteroidtreated patients) have a high risk of fatal infection associated with overwhelming infection not typically observed in immunocompetent patients.19 No deaths have been reported as a result of infection with E. ewingii.18,19 55.1.2.5  Pathogenesis of Human Ehrlichiosis The relatively low bacterial burden in the blood and tissues in nonimmunocompromised patients with HME suggests that the pathophysiology of ehrlichiosis may involve immunopathologic responses that are manifest as a toxic shock–like syndrome.3,119 The first murine model of fatal human ehrlichiosis has been instrumental in understanding the mechanisms behind the toxic shock–like syndrome of severe and fatal human ehrlichiosis.120 Mice inoculated with an ehrlichia (Ixodes ovatus ehrlichia; IOE) closely related to E. chaffeensis develop histopathology resembling that observed in HME patients, and a similar disease course is observed in the IOE murine model. Lethal infections with IOE are accompanied by extremely high levels of serum TNF-α,

TABLE 55.2 Clinical Manifestations in Published Series of HME Cases Clinical Feature

Eng et al.

Fishbein et al.

Everett et al.

Schutze et al.a

Olano et al.

Olano et al.

Fever Malaise/weakness Chills/Rigor Cephalea Myalgia Arhtralgia Anorexia Nausea Vomiting Diarrhea Diaphoresis Abdominal pain Cough Pharyngitis Rash Confusion Photophobia Stupor Coma Seizures Lymphadenopathy Jaundice Dyspnea Hepatomegaly Splenomegaly

100 61 65 77 53 28 50 54 49 38 21 23 39 33 47 29 11 NR NR NR 26 15 23 15 15

97 84 61 81 68 41 66 48 37 24 53 21 26 25 36 20 NR NR NR NR 25 NR NR NR NR

100 30 73 63 43 33 27 50 27 10 NR 10